BIOCATALYSIS RESEARCH PROGRESS No part of this digital document may be reproduced, stored in a retrieval system or transmitted in any form or by any means. The publisher has taken reasonable care in the preparation of this digital document, but makes no expressed or implied warranty of any kind and assumes no responsibility for any errors or omissions. No liability is assumed for incidental or consequential damages in connection with or arising out of information contained herein. This digital document is sold with the clear understanding that the publisher is not engaged in rendering legal, medical or any other professional services.
BIOCATALYSIS RESEARCH PROGRESS
FRANCESCO H. ROMANO AND ANDREA RUSSO EDITORS
Nova Biomedical Books New York
Copyright © 2008 by Nova Science Publishers, Inc.
All rights reserved. No part of this book may be reproduced, stored in a retrieval system or transmitted in any form or by any means: electronic, electrostatic, magnetic, tape, mechanical photocopying, recording or otherwise without the written permission of the Publisher. For permission to use material from this book please contact us: Telephone 631-231-7269; Fax 631-231-8175 Web Site: http://www.novapublishers.com NOTICE TO THE READER The Publisher has taken reasonable care in the preparation of this book, but makes no expressed or implied warranty of any kind and assumes no responsibility for any errors or omissions. No liability is assumed for incidental or consequential damages in connection with or arising out of information contained in this book. The Publisher shall not be liable for any special, consequential, or exemplary damages resulting, in whole or in part, from the readers’ use of, or reliance upon, this material. Independent verification should be sought for any data, advice or recommendations contained in this book. In addition, no responsibility is assumed by the publisher for any injury and/or damage to persons or property arising from any methods, products, instructions, ideas or otherwise contained in this publication. This publication is designed to provide accurate and authoritative information with regard to the subject matter covered herein. It is sold with the clear understanding that the Publisher is not engaged in rendering legal or any other professional services. If legal or any other expert assistance is required, the services of a competent person should be sought. FROM A DECLARATION OF PARTICIPANTS JOINTLY ADOPTED BY A COMMITTEE OF THE AMERICAN BAR ASSOCIATION AND A COMMITTEE OF PUBLISHERS. Library of Congress Cataloging-in-Publication Data Biocatalysis research progress/Francesco H. Romano and Andrea Russo(editors). p.;cm. Includes bibliographical references and index. ISBN 978-1-61668-287-3 (E-Book) 1.Enzymes—Biotechnology.2.Microbial biotechnology.I.Romano,Francesco Russo,Andrea,1954[DNLM:1. Catalysis.2. Enzymes.3.Biochemistry—methods. QU 135 B6147 2008] TP248.65.E59B568 2008 572’.—dc22 2008012763
Published by Nova Science Publishers, Inc.
H.,1939-II.
New York
Contents Preface
vii
Chapter I
Nanobiocatalytic Systems: Thin Films of Enzymes Laura Pastorino and Svetlana Erokhina
Chapter II
Bioprocesses for the Synthesis of Nucleosides and Nucleotides Marco Terreni, Daniela Ubiali, Teodora Bavaro, Davide A. Cecchini, Immacolata Serra and Massimo Pregnolato
Chapter III
Biocatalysis in Environmental Technology D. M. G.Freire, M. L. E. Gutarra, V. S. Ferreira-Leitão, M. A. Z. Coelho and M. C. Cammarota
Chapter IV
Application of Lipases to Substances with Pharmacological Importance (Polyunsaturated Fatty Acids) Marie Zarevúcka and Zdeněk Wimmer
Chapter V
The Evolution of Directed Evolution Birthe Borup, Lynne Gilson, Richard Fox and Thomas Daussmann
Chapter VI
Biocatalytic Resolution of DL-Pantolactone by Cross-linked Cells and its Industrial Application Zhi-Hao Sun, Ye Ni, Pu Zheng, Xin-Fu Guo and Jun Wang
Chapter VII
Biocatalytic Potential of Haloalkaliphilic Bacteria Satya P. Singh, Megha K. Purohit, Jignasha T. Thumar, Sandeep Pandey, Chirantan M. Raval and Hetal G. Bhimani
1
47
95
155 187
195
209
vi Chapter VIII
Chapter IX
Chapter X
Chapter XI
Chapter XII
Chapter XIII
Index
Contents Salt-tolerant Alkaliphilic Actinomycetes and their Biocatalytic Potential Satya P. Singh and Jignasha T. Thumar
219
Accelerating Whole-cell Biocatalysis by Cellular Membrane Engineering Ye Ni and Rachel R. Chen
229
Recent Advances in Enzymatic Synthesis of Water-Soluble Conducting Polymers Estibalitz Ochoteco, Tomasz Sikora, David Mecerreyes, Jose A. Pomposo and Hans Grande
245
Marine Enzymes for Biocatalysis – Production, Isolation and Applications K. Muffler, J. Mukherjee and R. Ulber
259
Immobilized Microbial Cells - Applications and Mass Transfer Phenomena Venko Beschkov
281
Application of Whole-cell Biocatalysis in Chemoenzymatic, Asymmetric Synthesis of Medically Important Compounds Ewa Żymańczyk-Duda and Paweł Kafarski
307 345
Preface Biocatalysis encompasses the use of enzymes or whole cell systems for effecting the conversion of readily available, inexpensive starting materials to high value products. Enzymes frequently display exquisite selectivity, particularly chemo-, enantio- and regioselectivity, making them attractive catalysts for a wide range of chemical transformations. Enzymes also typically operate under mild conditions of pH and temperature leading to the formation of products of high purity. As a result of these advantages, enzymes and whole cells are finding wider application in areas such as the production of intermediates for pharmaceuticals, fine chemicals, agrichemicals, novel materials, diagnostics, biofuels and performance chemicals. This new book presents leading-edge research in the field. Chapter I - The use of enzymes in industry requires their immobilization in order to reduce the costs and increase the yield of the biocatalytic processes. By reducing the enzyme mobility, the immobilization affects the enzyme structure and consequently its functionality. Thus, the immobilized enzymes efficiency is dependent on the peculiarities of the immobilization technique. These had been evolved from physical adsorption, membrane entrapment and chemical surface activation. Although these techniques are at present successfully utilized, the search for new approaches, which can result in biocatalytic systems with enhanced efficiency, is in progress. In particular traditional immobilization techniques do not provide effective control of the immobilization process or of the properties of the biocatalytic system at molecular level. This lack of control results in the formation of enzyme molecules aggregates leading to loss in functionality due to mass transfer resistance. This limitation can be overcame by nanotechnology inspired biocatalytic systems that differ from traditional systems in terms of preparation, catalytic efficiency and application potential. In particular the “bottom-up” approach, which consists of building systems molecule-by-molecule, represents a promising methodology for the precise control of the biocatalytic system structure. Moreover the molecular dimension of the resulting systems makes economically possible the use of even highly expensive material, including the most of enzyme molecules. The bottom-up approach comprises several methods, among them thin film techniques offer the possibility to construct 2 and 3D biocatalytic systems with nm resolution, with predetermined structure and function, starting from mono/multi layers of enzymes. This can be accomplished by the use of the layer by layer alternate adsorption of enzymes with oppositely charged polyions (layer-by-layer
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self assembly technique), or by the Langmuir-Blodgett technique based on the formation of enzyme-containing monolayers at the air/water interface. Enzyme layers can be deposited onto solid supports of different shapes or enzymes can be encapsulated into nano- and microcapsules with layered shell organization for the design of nano-bioreactors. Interest in this field is rapidly growing and is likely to fuel more exciting developments in the near future. The aim of this chapter is to review current status of layer-by-layer self assembly technique and of Langmuir-Blodgett technique for the development of biocatalytic systems, to give an overview of the techniques for the structural and functional characterization of enzyme thin films and to give some perspectives of future developments. Chapter II - Modified nucleosides and nucleotides are routinely used as citotoxic or antiviral drugs as well as immunosuppressive agents. The therapeutic activity of these compounds is due to their ability to act as antimetabolites in the RNA and DNA synthesis. Nucleosides have traditionally been prepared by various chemical methods, involving multistep chemical procedures which are plagued by low yields and the formation of undesired byproducts. These drawbacks strongly reduce the performances of the processes used for the synthesis of unnatural nucleosides and nucleotides, in terms of yields, purity of the final product, costs and environmental impact. Enzymatic syntheses have been shown to be an advantageous alternative to chemical methods due to the high selectivity of the enzymes, the mild reaction conditions and the overall simplicity of the approach. These bioprocesses can overcome the need of specific protecting groups often required on the heterocyclic base and/or on the sugar residue in the glycosylation reaction as well as in the modification of naturally occurring nucleosides. Consequently, these processes can be very competitive in terms of costs, allowing high quality products to be obtained in high yields. Glycosyl-transferring enzymes (e.g. nucleoside phosphorylases, Ndeoxyribosyltransferases) have been used in the synthesis of nucleosides by mediating the enantioselective transfer of glycosyl residues to acceptor bases. Recent reports involving glycosyl-transferring enzymes are reviewed herein. In this context, the use of isolated deaminases to obtain, respectively, cytidine derivatives from uridine, or guanosine derivatives from adenosine is also considered. Moreover, hydrolases (e.g. lipases, esterases) will be described in carrying out regio-controlled manipulations on the carbohydrate moiety. Finally, the enzyme-mediated synthesis of nucleotides through regioselective phosphorylation of the parent nucleosides catalyzed by kinases and phosphatases are discussed. This review will also focus on the importance of the enzyme selection and the design of the final biocatalyst thereof, particularly by tailor-made immobilization of the protein on solid support, with the aim to have an active, stable and reusable biocatalyst for preparative purposes. Chapter III - New strategies development to solve problems related to waste disposal may include technologies facing compounds of low biodegradability and/or detoxification of some residues with the alternative of adding value. There are various chemical and biological approaches, but biological ones employing enzymes had been showed good results. On the other hand, environmental use of enzymes depends on costs reduction for biocatalyst production through the selection of highly-productive strains, development and optimization
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of fermentation processes. Production system which employs agroindustrial residues as culture medium can be an interesting alternative. Microorganisms’growth on solids wastes by solid state fermentation can add value to these residues by enhancing nutritional quality and/or producing several enzymes with biotechnological applications, and also can reduce its pollution potential by degrading toxic compounds. The reduction cost of enzyme production could make feasible its employment for environmental purposes such as enzymatic pretreatment of wastewaters with high levels of fats, improving the biological degradation, or in the treatment of colored and phenolic wastes. The potential of environmental biocatalysis not only help to solve pollution problems but also simultaneously add value to undesirable wastes by the generation of biotechnological products or by enhancing its nutritional proprieties for use as animal feed. Chapter IV - Polyunsaturated fatty acids and their derivatives are extensively studied natural compounds with high impact in human medicine. Their sources belong among the renewable resources, mostly of plant origin. Lipases are well established biocatalysts for the enantio- and regioselective formation and hydrolysis of ester bonds in a wide variety of natural and unnatural substrates. Therefore, they seemed ideally suited also for bioconversion of the plant materials, especially plant oils. The use of biocatalysts for preparation of partial acylglycerols could provide numerous advantages as compared to conventional chemical methods such as increased selectivity, higher product purity and quality, energy conservation and the elimination of application of toxic catalysts. Two general routes to desired molecules are available, in principle namely hydrolysis/alcoholysis of triacylglycerols and esterification of glycerol. The reactions can be provided under conventional conditions or in supercritical fluids. Enzymatic reactions in supercritical fluids combine the advantages of biocatalysts (substrate specificity under mild reaction conditions) and supercritical fluids (high mass-transfer rate, easy separation of reaction products from the solvent, environmental benefits). Chapter V - The recent increasing usage of biocatalysts as viable alternatives to conventional chemical reactions is attributable to improved technology in tailoring enzymes found in nature to the needs of the industry. Arguably the most successful technology in this field has been directed evolution. Here the authors present the various stages in the evolution of directed evolution itself. The initial approach employed in directed evolution was successive rounds of random mutagenesis and screening, from which the top performer was chosen as parent for the next round of mutagenesis. Although effective, this method is not very efficient, as all beneficial mutations not found in the top performer are lost in the next round of evolution. With the advent of DNA recombination methods, mutations found in lower performing enzymes could be included in the next round. However, there was still useful diversity left unrecognized and unutilized, since only the top performing enzymes would be used for recombination. As statistical tools based on sequence-activity relationships were developed to help identify good mutations for recombination, beneficial diversity from even poorly performing enzymes could be moved to the next round of evolution. Presently, the drive to increase knowledge of sequence-activity relationships for enzyme families is enabling collections of variants to be synthesized that efficiently explore the chemical space of potential substrates for those enzymes. This will continue to lead to even faster evolution
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cycles, as a more relevant portion of the sequence space to be explored has been designed into the first library (first set of enzymes to be screened). Chapter VI - D-Pantoic acid (D-PA) and d-pantolactone (D-PL) are known as important intermediates for the production of calcium pantothenate, an additive for animal feed. Studies have showed that several filamentous fungi belonging to the genera Fusarium, Gibberella and Cylindrocarpon could produce D-PL hydrolases that selectively hydrolyze the D-isomer of DL-pantolactone to form D-PA,. In our previous studies, an excellent stereoselective D-lactonohydrolase producing strain Fusarium moniliforme CGMCC 0536 (SW-902) was obtained from isolation and mutation. Cells of this strain were successfully applied for the kinetic resolution of DL-pantolactone to produce D-(−)-isomer (99% e.e.). Furthermore, several immobilization methods were established to reuse the cells. The rationale for choosing whole cell immobilization is because it not only eliminates enzyme purification and extraction steps, but also increases enzyme thermostability, provides higher operational stability, greater resistance to environmental fluctuations and lower enzyme cost. An important technique is to treat cells with glutaraldehyde. The cross-linking of enzymes with glutaraldehyde involves the reaction between this bifunctional reagent and free amine groups of the enzyme. The linkages formed are irreversible and lead to cross-linked enzymes and cells exhibiting high operational stability. The wide utilization of glutaraldehyde for whole cells immobilization can be attributed to the decrease of enzyme leakage by cross-linking among the chemical reagent, cell wall, and intracellular protein. Therefore, the authors investigated the potential of crosslinking F. moniliforme CGMCC 0536 with glutaraldehyde for entrapping D-lactonohydrolase inside the cells and the employment of cross-linked biocatalyst in a stirred reactor. Their results have showed that cells of F. moniliforme CGMCC 0536 have been successfully immobilized by cross-linking with glutaraldehyde. The cross-linked cells exhibited a markedly improved thermal stability and operational stability than free cells. Kinetic characteristics of immobilized cells were assessed. The Km value of cross-linked cells (118 mM) was slightly higher than that of free cells (96 mM), while the Vmax value decreased from 4.18 to 3.87 mM min−1 g−1 wet cells after cross-linking. Furthermore, glutaraldehyde treatment did not change the stereospecificity, pH, and temperature profile of the D-pantonohydrolase. The high storage stability reduces fermentation workload. Significant process engineering advantages were evident for cross-linked cells in repeated batch operations. The resolution of DL-pantolactone was maintained at a steady level during 110 consecutive batches. The high activity and operational stability of the cross-linked cells presented in this work have been successfully implemented in the commercial production. Recently, the authors attempted to operate the resolution continuously by recycling cross-linked cells in a membrane bioreactor. The data show that glutaraldehyde cross-linking affords a satisfactory method for preserving the asymmetric hydrolyzing capacity of F. moniliforme CGMCC 0536. A feasible method with high operational stability for the production of D-PA catalyzed by cross-linked cells was established in a continuous process by using membrane bioreactor. Chapter VII - Haloalkaliphilic organisms have been largely investigated from Soda lakes around the globe and other habitats for these organisms are rarely explored. During the last 10 years, the authors have been working on the diversity and enzymatic potential of
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haloalkaliphilic bacteria from the natural and man made saline habitats along the coastal Gujarat in Western India. The organisms have displayed varied diversity based on their cultural and morphological patterns, Gram reaction, biochemical properties, antibiotic resistance-sensitivity, molecular phylogeny and secretion of extracellular enzymes. The production of extracellular alkaline proteases and lipases were widely spread among the isolates, whereas only few secreted amylase. The wide spread distribution of these bacteria from beyond soda lakes clearly indicated their ecological significance. Besides, the enzymatic potential would attract several biotechnological applications under alkalinity and high salt conditions. While major attention has been focused on molecular phylogeny and diversity, only limited information is available on their enzymatic potential and enzyme characterization. Cloning, sequencing and expression of different enzymes from Haloalkaliphilic bacteria and archaea are limited in literature. Our studies with alkaline proteases from a range of Haloalkaliphilic isolates revealed that many enzymes were highly resistant to urea denaturation and displayed catalytic potential under combination of extreme conditions. However, the ability to catalyse under extreme conditions was salt dependent. The enzymes displayed unique features for biotechnological applications, besides providing a model system to study protein folding and stability. It is evident that the secretion and properties of alkaline proteases would also be very useful in assessment of microbial heterogeneity. The prospects of metagenomics to explore the novel sequences for biocatalysts from noncultivable microbes have emerged as a potential tool in recent years. This would also be discussed with particular reference to saline habitats. Over all, our findings would be integrated with the literature and biocatalytic potential of Haloalkaliphilic bacteria and archaea would be assessed. Chapter VIII - Extremophiles are distributed over a range of extreme habitats. Among them, extremophilic actinomycetes have recently attracted greater attention due to their various natural products and specific mechanism of adapting extreme environments. The natural and man-made environments may harbor a large population of halophilic and alkaliphilic actinomycetes. However, they have only recently focused attention of the researchers. The phylogeny, diversity and biotechnological potential of salt-tolerant alkaliphilic actinomycetes are still in infancy. It is, therefore, relevant and important to pay more attention to extreme actinomycetes from unexplored habitats, as a possible way to discover novel taxa and, consequently, new secondary metabolites. The study would also enlighten us on their diversity, phylogeny and ecological significance. During the last decade, there has been a dramatic increase in the need for bioactive compounds with novel activities. Enzymes, after antibiotics are the most important biologically derived product having immense potential in catalytic reactions of commercial interest. Most of the studies related to enzymes have so far focused on halophiles, alkaliphiles and haloalkaliphiles; however, the enzymatic potential of halo-tolerant alkaliphilic actinomycete is nearly untouched. From the literature, it is evident that the exploration of the enzymatic potential of these microbes is just the beginning and till date only few enzymes are investigated in depth. Salt-tolerant alkaliphilic actinomycetes produce enzymes, such as alkaline protease, amylase, cellulase and lipase that are functional under extreme conditions. Consequently, the unique properties of these biocatalysts have potentials in several novel
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applications in industrial processes. Because of the capacity to survive under nonstandard conditions in non-conventional environments, it is assumed that the properties of extremophilic enzymes have been optimized for these conditions. Interestingly, our own studies have also revealed the occurrence of alkaline protease, amylase and cellulase in salttolerant alkaliphilic actinomycetes isolated from vast coastal line and saline habitats along the Saurashtra Coast Gujarat (Western India). In addition, the results on the alkaline proteases are suggestive of their unique position in the generation of novel enzymes. Chapter IX - Whole-cell biocatalysts are preferred in biocatalysis applications involving cofactors and/or multiple enzymes. However, cell envelope often represents a formidable permeability barrier, limiting the rate of entry of substrate. As a result, reactions catalyzed by whole-cells are reportedly orders of magnitude slower than those of by their free enzyme counterparts. This chapter reviews recent molecular engineering efforts addressing this critical issue. Initial studies were carried out with E. coli strains carrying mutations in the outer membrane structures. The effects of these mutations were investigated by interrogating the mutant cells with substrates differing substantially in size and hydrophobicity. The reduction of outer membrane permeability barrier by these mutations led to significant accelerations in reaction rates of all whole-cell catalyzed reactions investigated. In the case of the tetrapeptide, a substrate for subtilisin, a single gene mutation in lipopolysaccharide (LPS) synthesis can render the outer membrane completely permeable to substrate, reaching a barrier-less condition that maximizes the reaction rate while retaining all the benefits of whole-cell catalysts. For reaction rates of toluene dioxygenase (TDO)-catalyzed reactions, an increase of up to 6-fold was observed with lipoprotein mutant for three small, hydrophobic substrates tested. Mutations in either LPS or lipoprotein are effective for accelerating reactions with UDP-glucose, a hydrophilic molecule with Mw over 600 Da, resulting in a striking acceleration (up to 14-fold) of reaction rate. The magnitude of reaction rate acceleration was found to be dependent upon the substrate concentrations, the enzyme expression level, and the nature of the mutations. In addition, the mutations were demonstrated to be far more superior to common permeabilization procedures such as freezethaw and EDTA treatments. To understand the mechanism of the permeability enhancement in the lipoprotein mutant, the lpp region was sequenced. The results revealed that Braun’s lipoprotein was absent in the transposon mutant cells due to multiple stop codons within the 59-bp insertion, suggesting that the absence, rather than the alteration of Lpp, is responsible for the observed change in permeability. More importantly, the sequencing result suggests that lpp deletion could become a general permeabilization method. Subsequent studies were carried out by generating lpp deletion mutants from strains with different genetic background. It was indeed shown that lpp deletion generates a useful phenotype, not only effective in enhancing substrate permeability, but also in reactions limited by product permeability, as demonstrated in L-carnitine synthesis. Importantly, the deletion has no significant effect on cell growth, metabolism and recombinant protein expression. Therefore, lpp deletion phenotype could be applied as a generally applicable method to enhance the outer membrane permeability of various E. coli strains, and possibly other Gram-negative bacteria with lpp homologs.
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Chapter X - The present chapter introduces the newest synthesis strategies developed by our research group in order to overcome the main disadvantages derived from enzymatic biocatalysis as a strategy for producing conducting polymers. First, the enzymatically catalyzed polymerization of 3,4-ethylenedioxythiophene (EDOT) in the presence of polystyrenesulfonate (PSS) is achieved showing an electrical conductivity of 2×10-3 S/cm and an excellent film formation ability as confirmed by atomic force microscopy images. Second, a simple method for immobilizing horseradish peroxidases (HRP) in the biocatalytic synthesis of polyaniline (PANI) is presented. This method is based on a biphasic catalytic system where the enzyme is encapsulated inside the ionic liquide (IL) 1-butyl-3methylimidazolium hexafluorophosphate, while other components remain in the aqueous phase. The enzyme is easily recovered after reaction and reused several times. Finally, a new bifunctional template (sodium dodecyl diphenyloxide disulphonate, DODD) is proposed in the synthesis of polyaniline (PANI) as a strategy to improve water solubility as well as electrical conductivity in the obtained polymer. Chapter XI - The diversity of marine life offers a variety of novel enzymes which might have a tremendous potential as biocatalysts for academic research as well as for industrial processes. Concerning the wide spectrum of ecological habitats, which differs extremely in temperature, pressure, and salinity, the whole machinery of enzyme expression and stability of the proteins was adopted to the distinct environmental conditions of the producing microas well as macroorganisms. Therefore the oceans provide an almost untapped reservoir of biocatalysts showing interesting properties like high salt, pressure, and temperature tolerance. With respect to the special requirements of industrial processes which are particularly focused on high mass transfer and high time space yields, respectively, enzymes from marine origin located at exotically regions can help to fulfill these specific demands. Thus, applications of such proteins or whole cells as catalysts for manufacturing bulk- and fine chemicals seems visionary up to know. However, regarding the current trends and developments in molecular biology as well as genetic engineering it appears more realistic in the case of long-term view. By means of identifying the origin of the enzymes’ stability one has the possibility to modulate other so far unstable (terrestrial) enzymes. This review covers the development and research work done on the processing of enzymes from marine origin, which can be used as biocatalyst tools for research in academia and industry. Chapter XII - Immobilized cells performance has been extensively studied in the last three decades. The advantages of this type of biocatalysts are in their multiple use, continuous operation and easy removal from the reaction mixture. Two main types of immobilization techniques exist: cell entrapment in gels and cell fixation on solid supports. The drawbacks of cell entrapment are in the diffusion limitations at substrate supply to and product removal from the gel particles. Such limitations do not exist in cases of cell fixation on solid supports but other transport phenomena arise, associated with the cell accumulation and concentration within small area. Both techniques admit changes in the cell physiology including microbial growth different from that in free culture. That is why additional impact on the net biocatalyst performance may cause the microbial growth in immobilized state and the possible cell leakage into the fermentation broth and their consecutive growth in a free state.
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The choice of immobilization method depends on the kinetics of the very microbial process, on the microbial growth, on the products of reactions, being possible inhibitors or catalysts, etc. The purpose of this article is to review and to discuss the advantages and the drawbacks of these two groups of immobilized biocatalysts from the point of view of the specific reaction kinetics and the mass transfer limitations. Mathematical modeling is employed to evaluate the effects of cell growth and detachment from the particles and their contribution in free and immobilized state. Experimental data are presented to illustrate the discussed effects. Chapter XIII - Chirality is, in the most cases, the key factor in the safety and efficacy of many drug products. Usually only one enantiomer is responsible for the desired activity, whereas its counterpart could be inactive, possess some activity of interest, be an antagonist of the active enantiomer or have a separate activity that could be either desirable or undesirable. Only in a few cases specific compositions of a racemate or an enantiomeric pair demonstrated a synergistic effect. In past decades the pharmacopoeia was dominated by racemates, but since 1980 number of chiral drugs introduced to market have grown markedly. Chiral compounds currently account for at least 50% of sales with the annual sales of singleenantiomer drugs exceeding 150 billions of dollars in 2002. These compounds now represent one-third of all drug sales worldwide. Thus, it is not surprising that chiral intermediates and fine chemicals are in high demand, both from the pharmaceutical and agrochemical industries. During the drug development process, the question invariably arises of which step and which method to choose for introducing chirality. Thus, racemates are usually produced by parallel synthesis for drug candidates - it makes sense to resolve the racemate for the initial animal studies, whereas an optimized, enantioselective synthesis is employed in the course of production. While enantiomerically selective organic synthesis is the traditional approach, using enzymes and enzyme-containing microorganisms to biocatalyze a reaction is becoming increasingly important. However, only 85% of products obtained by biocatalytic processes are enantiomerically pure and in 50% of these processes enantiomerically active substrates are used. This shows that induction of asymmetry is not always the most important feature of industrial biocatalysis. Quite successfully it can be used to carry out conversions that would otherwise require difficult or multiple synthetic chemistry steps. In such cases, biocatalysis can be the preferred route even if chirality is not desired. Moreover, over 25% processes base on kinetic resolution, reaction which affords the desired products with maximal yield of 50% and thus seems to be not interesting from industrial point of view. This is because the second enantiomer is either useful (although in different processes) or is isomerized and processing back as subtrate. The microbial biocatalysts demonstrate a wide variation of activities between genera, within genera and even within species. The range of substrates transformed and the inter- and intra-species differences in specificity of the individual biocatalysts suggests, that it is possible to provide multiple catalytic agents, especially when the traditional organic process fails or is to expensive to perform. Thus, biocatalysis have become an attractive alternative to conventional catalysts in numerous industrial processes. When compared to chemical catalysis biocatalysts are exquisitely selective and highly precise due to: their substrate selectivity, which allowed distinguishing and acting on the subset of compounds within a
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larger group of chemically related compounds; their stereoselectivity - the ability to act on a single enantiomer or diastereoisomer selectively and their regioselectivity – ability to recognize one location in a molecule and finally because of their selectivity towards defined functional group in a presence of other equally reactive or more reactive ones. Furthermore, biocatalysts are able to carry out bioconversions under mild conditions – another benefit of using them as industrial catalysts. Biocatalysis in drug reseach and production is usually used in multistep processes in concert with more traditional production techniques. Thus, biocatalysts (as both isolated enzyme and whole-cell systems) are increasingly being used to assist in synthetic routes to complex molecules of industrial interest, in so called chemoenzymatic processes. Whole-cell biocatalysis is a useful alternative to the use of pure enzymes. Employing whole – cells is a strategy which allowed overcoming many limitation of the enzymatic biotransformations and, what is important, is usually cheaper than using purified enzymes. Additionally, microbial cells used as a catalysts, have their own cofactor regeneration systems, offer wide range of enzymatic activities towards a number of non-physiological substrates and moreover, usually there are no side reactions except the expected ones. Moreover, there is no doubt that, the advantage of microbial biotransformations is the possibility to induce enzymes of defined, desired activity even if they are not constitutively presented inside the microbe cells. This enlarge the offer of possible reactions to be carried out and in fact there exists an enzyme for almost every type of chemical reaction.
In: Biocatalysis Research Progress Editors: F. H. Romano, A. Russo
ISBN: 978-1-60456-619-2 © 2008 Nova Science Publishers, Inc.
Chapter I
Nanobiocatalytic Systems: Thin Films of Enzymes Laura Pastorino1 and Svetlana Erokhina2 1
Department of Communication, Computer and System Sciences, University of Genova, Italy 2 Department of Physics, University of Parma, Italy
Abstract The use of enzymes in industry requires their immobilization in order to reduce the costs and increase the yield of the biocatalytic processes. By reducing the enzyme mobility, the immobilization affects the enzyme structure and consequently its functionality. Thus, the immobilized enzymes efficiency is dependent on the peculiarities of the immobilization technique. These had been evolved from physical adsorption, membrane entrapment and chemical surface activation. Although these techniques are at present successfully utilized, the search for new approaches, which can result in biocatalytic systems with enhanced efficiency, is in progress. In particular traditional immobilization techniques do not provide effective control of the immobilization process or of the properties of the biocatalytic system at molecular level. This lack of control results in the formation of enzyme molecules aggregates leading to loss in functionality due to mass transfer resistance. This limitation can be overcame by nanotechnology inspired biocatalytic systems that differ from traditional systems in terms of preparation, catalytic efficiency and application potential. In particular the “bottom-up” approach, which consists of building systems molecule-by-molecule, represents a promising methodology for the precise control of the biocatalytic system structure. Moreover the molecular dimension of the resulting systems makes economically possible the use of even highly expensive material, including the most of enzyme molecules. The bottom-up approach comprises several methods, among them thin film techniques offer the possibility to construct 2 and
2
Laura Pastorino and Svetlana Erokhina 3D biocatalytic systems with nm resolution, with predetermined structure and function, starting from mono/multi layers of enzymes. This can be accomplished by the use of the layer by layer alternate adsorption of enzymes with oppositely charged polyions (layerby-layer self assembly technique), or by the Langmuir-Blodgett technique based on the formation of enzyme-containing monolayers at the air/water interface. Enzyme layers can be deposited onto solid supports of different shapes or enzymes can be encapsulated into nano- and microcapsules with layered shell organization for the design of nanobioreactors. Interest in this field is rapidly growing and is likely to fuel more exciting developments in the near future. The aim of this chapter is to review current status of layer-by-layer self assembly technique and of Langmuir-Blodgett technique for the development of biocatalytic systems, to give an overview of the techniques for the structural and functional characterization of enzyme thin films and to give some perspectives of future developments.
Introduction Enzymes molecules are catalysts and they speed up the chemical reactions at the basis of the metabolism of all living organisms with extremely high selectivity and efficiency. Moreover, they are not part of the final product of the catalysed reaction, and they operate at mild conditions, such as room temperature and physiological pH [Hartmeier, 1988]. In this respect enzymes can be considered remarkable catalysts and their catalytic properties represent the environmentally friendly solution to industrial problems. From the application point of view, it is very important that many enzymes are commercially available, and numerous industrial applications have been described. Enzymes have been used for more than 50 years in the detergent, textile, food and feed industries [van Beilen, 2002]. Moreover, biocatalysis has an important role in the industrial synthesis of bulk chemicals [Koeller, 2001], pharmaceutical [Pollard, 2006] and agrochemical intermediates [Aleu, 2006]. Biocatalysis also holds considerable promises in environmental fields [Alcade, 2006], such as enzymatic bioremediation [Whiteley, 2006; Wu, 2008], enzyme-based biofuel cells [Kim, 2006a] and biodiesel production [Rashid, 2008; Hernández-Martín, 2008]. These are only few examples of the growing impact of biocatalysis on different industrial sectors. However, the potentiality of enzymes in industrial applications is not exhaustively exploited, because of some bottlenecks such as the high cost of the most of enzymes, the low activity and/or stability under working conditions and low reaction yields [Coward-Kelly, 2006]. Recent advances in tools and techniques for biocatalysis research and development should help to overcome these limitations [Bommarus, 2006; Coward-Kelly, 2006; van Beilen, 2002]. Advances in metabolic and protein engineering, high-throughput screening, nanotechnology and other technologies will increase the impact of biocatalysis in industry [Rubin-Pitel, 2006; Coward-Kelly, 2006] and new industrial applications are expected to be realized [Schmid, 2001; Schoemaker, 2003]. Different steps can be individuated in the development of a biocatalytic process. They span from the identification of the target reaction, to the selection of the more suitable biocatalyst, to its characterization and modification, in order to arrive finally to its application
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[Schmid, 2001]. The economic feasibility of a biocatalytic process depends on several factors involved in these steps and differs for each process under study since requirements vary enormously depending on the biocatalyst itself. One critical point to be taken into account is represented by the cost of the biocatalyst itself, as many enzymes are expensive. To minimize the influence of this cost on the whole process, recombinant DNA technologies [Petersen, 1999; Sheldon, 2004] can be used to enable the production of enzymes with competitive prices on the one hand and immobilization techniques make possible the reuse of the same biocatalyst in different production cycles on the other hand [Hartmaier, 1988; Schmid, 2001; van Beilen, 2002]. As relates to immobilization techniques, they represent a powerful tool not only to reduce the cost related to the enzyme itself but also to simplify the design of the production plant, as the reaction product can be easily recovered without enzyme contamination, to simplify the process control and finally to improve some enzyme properties such as operational stability [Katchalsky-Katzir, 1993; Mateo, 2007]. Depending on the process requirements (enzyme characteristics, substrate, reaction type and reactor configuration) specific immobilized enzymes should be designed. Moreover, several possible restrictions, such as activity losses and diffusional limitations, should be taken into account in the design of immobilized enzymes [Cao, 2005]. For these reasons different immobilization strategies have been set up so far. They can be divided into five main groups based on the kind of interactions between the enzyme molecules and the solid support, and specifically they are: adsorption, micro-encapsulation, entrapment, cross-linking and covalent bonding [Hartmaier, 1988; Chibata, 1986, Katchalski-Katzir, 1993]. Even if many strategies are already available and have also found some applications at the industrial scale, the search for innovative and more efficient approaches for the immobilization of enzymes is still under progress [Cao, 2005]. One of the main limitation of traditional approaches is the lack of the effective control of the immobilization process at the molecular level. This lack of control results mainly in the formation of aggregates of enzyme molecules, which have partly lost their active conformation, in which the active sites could be not exposed to the substrate molecules and where the catalytic activity is lowered also by mass transfer resistance. It is then evident how the development of immobilization techniques able to control the positioning and the orientation of the enzyme molecules is required in order to control finally the properties of the realized biocatalytic medium. In that sense, nanotechnology appears to have a pivotal role in the development of the next generation of biocatalysts. Nanotechnology has found applications in almost all fields of research from aerospace to electronics, biology, medicine and biotechnology. Recently in the field of biotechnology a new area has appeared, which is nanoscale enzymatic biocatalysis [Wu, 2004]. The area of enzymatic nano-biocatalysis comprises both the use of nanostructured materials as hosts for enzyme immobilization [Kim, 2006b; Wang 2006], via approaches including adsorption, covalent attachment and so on, and the fabrication of biocatalytic systems molecule-bymolecule with a predetermined structure and, therefore, function [Huie, 2003; Schäffer, 2007]. This last approach to nanofabrication is the so called “bottom-up” one and is based on the use of chemical or physical forces at the nanoscale level to assemble molecules, acting as building blocks, into complex structures with a predetermined architecture and function [Shimomura, 2001; Seeman, 2005; Zhou, 2005]. Extensive research has been performed on
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the bottom-up approach in the last years and a number of bottom-up methods have been developed also due to the appearance of techniques allowing a direct characterization at the molecular scale. Among the bottom-up nanofabrication methods, thin film techniques represent a powerful tool for the assembling of molecules into highly ordered architectures [Ulman, 1991; Seeman, 2005]. In particular thin film techniques are well suited for the manipulation of enzymes which display the capability to form highly packed assemblies and to generate 2D and 3D structures. The properties of the developed structures can be tailored by incorporating appropriate molecules and functional groups. There are different mechanisms by which thin films can be accomplished, two of them are the Langmuir-Blodgett technique (LB) and the layer-by-layer self assembly (LbL) technique. The LB technique, which was developed at the beginning of the twentieth century [Blodgett, 1935; Blodgett, 1937], is based on the formation of a closely packed monolayer at the air/water interface and on it subsequent transfer onto the surface of a solid support. This technique was mainly developed for the deposition of amphiphilic molecules. However, extensive work has been carried out on the formation of thin films of biological interest including enzyme molecules for biocatalytic purposes. The LbL technique, a more recently developed technique, is based on the alternate assembly of oppositely charged polyelectrolytes for the deposition of complex multilayered nanostructures [Iler, 1966; Decher, 1997]. This process is a very simple one and can be used for the deposition of multilayered films onto supports of any shape with the precise control of the structure and of the function. Enzyme molecules can be assembled using this technique for the fabrication of biocatalytic media or can be entrapped into nanocapsules for the fabrication of nanobioreactors. Variations of film composition and sequence of layer alternation influence the properties of the biocatalytic film. Moreover sequential catalytic reactions can be realized in multi-enzymes films. Enzymes immobilized in thin films have shown to posses unique properties in terms of stability and functionality. Therefore, these new types of biocatalytic systems would have a great impact on the industrial application of enzymes. The main aim of this chapter is to overview some of the more advanced and promising nanotechnological methods for the immobilization of enzymes and specifically the fabrication of organized thin film architectures by the LB technique and the LbL self assembly technique. The current status and applicability of these methods in the biocatalysis field, as well as prospective applications and developments are discussed.
The Langmuir-Blodgett Technique LB Introduction Langmuir-Blodgett (LB) technique is one of the most powerful tools for the realization of functional organic structures with molecular resolution in one direction. In particular, it was widely used for realization of layers with biological molecules. Lipids and lipid-like molecules are traditional objects of the applicability of the method. Proteins were also deposited using LB technique. However, in the case of proteins, the method not always can
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be applied directly for the deposition of protein-containing layers and, therefore, must be modified according to the nature of each particular protein. Strictly speaking, monolayers of only membrane proteins can be directly formed at the air/water interface. For all other proteins, including enzymes, special tools and approaches must be developed in order to form functionally active molecules. Reviews on the approaches allowing to overcome the difficulties can be found in [Erokhin, 2000; Erokhin, 2002]. Enzymes are proteins, whose function is to perform the activity of the extremely effective biological catalysers. Therefore, mutual orientation of the active zones in the enzyme molecule, responsible for the electron transfer from and to biological molecules (substrates), involved into the reaction, is very important for their functional activity. This feature is crucial considering the applicability of LB method for the enzyme layers realization. Essential step of the LB technique is spreading and compression of the layer at the air/water interface. This step is very important for all the protein monolayers in general, and even more critical for enzymes in particular. Exposition to the action of the surface tension of the pure water surface (72 mN/m) can result in the complete denaturation or at least partial modification of tertiary and/or secondary structure of the enzyme. In fact, The structure of the enzyme globule is stabilized by the molecular interactions within hydrophobic nucleus of the protein, that is comparable with the surface tension. These modifications of the protein structure can result in the irreversible decrease of the enzymatic activity. However, LB technique is still widely used for the working with enzyme molecules. It is worth to mention that the very first work on the application of the LB technique for enzyme layers formation was carried out in 1938 by I. Langmuir and V. Schaefer [Langmuir, 1938]. This work is well-known as it describe the first application of the horizontal deposition method (Langmuir-Schaefer technique). However, we would like to recall the fact (rarely mentioned now), that the work was performed on pepsin and urease monolayers.
Number of articles / year
20 18 16 14 12 10 8 6 4 2 0 1938
1948
1958
1968
1978
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2008
Year
Figure 1. Number of articles on enzyme-containing Langmuir and LB films per year (the dependence was averaged for 5 years).
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Literature analysis allows to monitor the genesis of the activity in the field of the application of LB technique to study properties of enzymes and/or deposition of enzymecontaining layers. Averaged dependence of the number of articles concerning the enzymecontaining Langmuir and LB layers on the year of publication is presented in Figure 1. Averaging was performed for 5 years in order to smooth occasional scattering and to reveal existing tendencies. Let us analyse the figure. First publication of I. Langmuir and V. Schaefer had not attract the attention of researched. Only one other article was published during almost 3 decades. However, at the end of the sixties we can see an increasing activities in the field coming to the maximum at the end of seventies. These activities can be connected to the fundamental investigations of the action of enzymes on the model membranes. In fact, Langmuir monolayer at the air/water interface can be consider as a model of the half of the biological membrane. Its composition can be varied by the variation of spreading solution using natural mixtures of lipids and/or mono- or multi-component solutions of different amphiphilic molecules. The state of the model membrane can be also varied by changing its surface pressure, modelling, therefore, liquid-expanded, liquid-condensed or solid regions of the biological membrane. Enzyme molecules were placed into the subphase under the monolayer and its activity was analysed by all available methods (in the early stages of the research, restricted number of techniques was available; the most of works were performed by the analysis of the monolayer area and surface potential variations). As it is practically always in the scientific activity, after the peak we can observe some decrease of the activity coming to the steady state situation. LB technique was found to be rather useful for the investigation and understanding of interaction of enzyme molecules with model membranes and such kind of works are still performed by different research groups. At the end of eighties we can observe a significant increase of the activity in the field of enzyme-containing LB films coming to the maximum in the beginning of nineties. This period coincides perfectly with wide spreading of the ideas of molecular and bio-electronics, as well as the beginning of the numerous works on the biosensors construction. This stage of researches are mainly dedicated to the incorporation of enzymes in thin layers and their transfer onto the surface of adequate transducer, capable to register the enzymatic reaction and, therefore, the presence of the substrate molecules in the solution under investigation. As in the previous case, some decrease of the activity was observed in the second part of nineties and we can see currently rather steady state situation, indicating continuous interest of research groups to the application of LB method for the construction of enzyme-containing functional molecular structures. The part of the work, dedicated to the Langmuir and LB films techniques for study the activity and construction of enzyme-containing layers, will be organized in the following manner. First, we will briefly present the basic principle of the LB technique in its classic form. Then, we will illustrate the applicability of the technique for the investigation of the interaction of enzymes with model membranes considering some examples, described in literature. Third, we will describe several special modifications of LB technique, allowing deposition of enzyme layers with preserved activity. It will be illustrated by some special cases of different enzymes. We will also present some examples of biosensors, using enzyme-
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containing LB layers as active sensitive elements. Finally, we will present a list of enzymes, ever used for LB studies, in order to provide reference points for further researches.
Basic Principles of Langmuir-Blodgett Technique Monolayers at the Air/Water Interface Salts of fatty acids are “classic” objects of the LB technique [Blodgett, 1937; Gaines, 1956; Overbeck, 1993; Peterson, 1992]. General structure of fatty acid molecules is:
CH 3 (CH2 ) n COOH
Surface pressure (mN/m)
Fatty acids, which form stable monolayers are stearic (n=16) one, arachidic (n=18) one, and behenic (n=20) one. Being place at the air/water interface, these molecules arrange themselves in such a way, that its hydrophilic part (COOH) penetrates water due to its electrostatic interactions with water molecules, which can be considered as electric dipoles or, more frequently, as electric charge, as head-group can be in a dissociated form. Hydrophobic part (aliphatic chain) faces itself to air, because it can not penetrate water due to entropy reasons. Therefore, if rather few molecules of such type were placed to the water surface, they form 2-dimensional molecular system at the air/water interface.
60 50 40 30 20 10 0 0
10
20
30
40
50
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80
Area per molecule A (A²)
Figure 2. π - A isotherm of stearic acid monolayer at the air-water interface.
Let us consider what will happen when the layer is compressed with some kind of barrier. We will consider surface pressure as a parameter describing the monolayer state. Surface pressure is determined as:
π = σ H O − σ ml 2
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where σH 0 is the surface tension of the pure water and σml is the surface tension of the 2 monolayer covered water surface. In other words, surface pressure can be considered as the decrease of the water surface tension due to the presence of the monolayer on it. Compression isotherm of the stearic acid monolayer is presented in Figure 2. This important characteristics represents the dependence of the surface pressure upon the area per one molecule, obtained at constant temperature. Usually, this dependence is called π-A isotherm. The measurements of the isotherm are practically always performed for studying the behaviour and phase transitions of the monolayers at the air/water interface. Let us consider the isotherm. Initially, the compression does not result in the surface pressure variations. Molecules at the air/water interface are rather fare from each other and do not interact. In some cases they also form domains, containing several molecules. This state of the monolayer is refereed as “two-dimension gas”. Further compression results in the increase of the surface pressure. Molecules come closer one to the other and begin to interact. This state of the monolayer is referred as “two-dimensional liquid”. For some compounds it is possible to distinguish also liquid-expanded and liquid-condensed phases. Continuation of the compression results in the appearance of “two-dimensional solid state” phase, characterized by the sharp increase in the surface pressure even for small decrease in the area per molecule. Dense packing of molecules in the monolayer is reached in this case. Further compression results in the collapse of the monolayer. 2-D structure does not exist anymore. Not controllable multilayers are formed at the water surface. Two instruments are usually considered for surface pressure measurements, namely, Langmuir balance [Langmuir,1917] and Wilhelmy balance [Wilhelmy,1863]. Langmuir balance measuring principle is the following. Barrier, separating the clean water surface from that covered with the monolayer, is the sensitive element. Even if this balance measures directly the surface pressure, it is not so frequently used. Mainly it is used when precise measurements of the monolayer conditions at the air/water interface are studied, and it is practically not used when the monolayer is supposed to be transferred onto solid substrates. There are several reasons for its restricted applications. First of all, the utilization of the Langmuir balance supposes that the compression of the monolayer is from one direction only. This fact can result in the gradient of the monolayer density, what is absolutely undesirable in particular cases. Second, the measurement of the surface pressure is performed in the other point with respect to that, where the deposition takes place. This feature can result in the weak control of the monolayer state during its transfer onto solid substrates. Third, there is rather large area of the monolayer, which cannot be used. The first and the second drawbacks are very critical when working with rigid monolayers. It was shown that it is possible to observe a gradient of the monolayer density (and so the surface pressure), if the compression is anisotropic one. The third drawback is very important when working with expansive substances, such as biomolecules (including enzymes), as a significant part of the layer must be wasted. Wilhelmy balance had found more applications, even if it does not provide the direct measurement of the surface pressure. However, it allows to avoid all 3 drawbacks of the Langmuir balance mentioned above. The sensitive element of the Wilhelmy balance is a plate (very often it is made from paper). The measurement principle is illustrated in the Figure 3.
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The forces acting to the plate are: mg -weight of the plate; FA - Archimedic force; Fs - surface tension induced force. The last force (FS) is just the product of the surface tension and the plate perimeter. As the weight of the plate is constant, and the Wilhelmy balances now are equipped by systems, maintaining the plate immersion depth into the water at the same level, providing, therefore, the Archimedic force constant, it is possible to attribute the zero value to the clean water surface, and the differences from this value will be directly the surface pressure. The construction of the balance allows to perform measurements in the point exactly corresponding to the deposition point with respect to the barrier position. It provides the precise control of the surface pressure value, maintained by the feedback system. The other advantage of the Wilhelmy balance is the possibility of compressing the monolayer from both sides, performing better homogeneity of the monolayer.
Figure 3. Illustration of the working principle of the Wilhelmy balance.
Figure 4. Typical dependences of the surface pressure (a) and surface potential (b) on the barrier coordinate for the monolayer at the air/water interface.
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As it was mentioned above, an important aspect of the Wilhelmy balance utilization is connected to the necessity of the maintenance constant of the plate position with respect to the water surface level, in order to avoid the variations in the Archimedic force. Usually, it is reached by special construction of the balance. Wilhelmy plate is connected to the magnet, which is inserted into the solenoid. Variations in the surface pressure displace the magnet position, and electronics provides the current in solenoid, which brings back the initial magnet position. The position is usually controlled by optical methods. The value of the current is proportional to the surface pressure value. Mentioned advantages allow to state that the most of measurements of the surface pressure are currently performed with Wilhelmy balance. The other parameter, which can be controlled when working with monolayers at the air/water interface, is the surface potential. The surface potential appearance results from the orientation of molecular charges and dipoles during the compression of the monolayer. Three different parts of the monolayer are usually considered for the surface potential interpretation [Oliveira, 1992]. The first one is due to the orientation of the C-H bonds in the hydrocarbon chains of the amphiphilic molecules during the monolayer formation. The second one is connected to the regular arrangement of the polar head-groups. And the third one is due to the dipol orientation of water molecules in the area just under the monolayer. The relative input of each regions can be different and it is due to the nature of the molecules forming the monolayer. Typical dependence of the surface potential of the monolayer upon the barrier coordinate is presented in Figure 4 [Yasutake, 1996] (the dependence of the surface pressure is presented in the same figure for the comparison). It is interesting to note, that there are differences in the behavior of the surface potential with respect to that of the surface pressure. The variation of the surface potential begins much before than the surface pressure begins to increase significantly. Such behavior is due to the fact, that molecules begin to aggregate, forming dimers, trimers and small domains, at the initial stage of the monolayer formation. Being aggregated, molecules tend to orient themselves in energetically adequate position, giving rise to the variation in the surface potential. It happens when there is practically no increase of the surface pressure. In the latter stage of the monolayer compression the variation of the surface potential is mainly due to only the increase of the monolayer density. Kelvin probe is the tool, which is usually used for the surface potential measurements [Surplice, 1970; Yasutake, 1996; Di Natale, 1999]. The instrument is equipped with a vibrating electrode, placed near the water surface. The reference electrode is inserted into the water subphase. Vibrating electrode and water surface form a capacitor. Vibration of the electrode provides the modulation of the capacity, resulting, therefore, in the appearance of the alternating current proportional to the value of the surface potential. The value of the surface potential of the monolayer can be both positive and negative and the sign of the surface potential is determined by the nature of the molecules in the monolayer.
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Monolayer Transfer onto Solid Substrates The floating monolayer can be transferred onto the surface of solid supports. Two main techniques are usually considered for the monolayer deposition, namely, Langmuir-Blodgett (or vertical lift) one [Blodgett, 1934] and Langmuir-Schaefer (or horizontal lift) one [Langmuir, 1938]. The scheme of the Langmuir-Blodgett deposition is illustrated in Figure 5. Specially prepared substrate with hydrophobic surface is passing vertically through the monolayer. The monolayer is transferred onto the substrate surface during this passing. The important point of such deposition is connected to the necessity of having the monolayer in the electrically neutral state. If some charges in the monolayer molecule head-groups will be uncompensated, the deposition will not be performed – electrostatic interaction of these charges with water molecules will be higher with respect to the hydrophobic interactions of their chains with the hydrophobized substrate surface. It will make impossible the monolayer transfer to the solid substrates. Let us consider again the monolayer of fatty acids in order to demonstrate the necessity of the head group neutrality. If the monolayer is formed at the surface of distilled water (pH is about 6.0) it cannot be transferred onto solid substrate. Its head group is dissociated and contains negative charge (COO-). There are two ways to provide the possibility of deposition. In the first case one must use LS technique (horizontal lift) and to provide the conditions when all head groups of the fatty acid molecules are in the protonated (not dissociated) form [Erokhin, 2000]. It requires the decrease of the pH of the subphase. In fact, the deposition begins to take place when the pH value is less then 4.0 when there is practically no dissociation.. However, the monolayer of pure fatty acids is very rigid and its transfer results usually in defective LB films at solid substrates. Therefore, usually fatty acid salt monolayers are deposited instead of pure fatty acids [Agarwal, 1973; Alekseev, 1987; Amador, 1998]. In this case, bivalent metal ions are added into the water subphase. Normally, their concentration is of the order of magnitude of 10-4 M. The metal ions attach themselves electrostatically to the dissociated fatty acid head groups, providing their electric neutrality. At the air/water interface, these ions are in a dynamic equilibrium with the fatty acid groups in the monolayer. In the deposited layer, the bivalent metal ion coordinates 4 oxygen atoms in two fatty acid molecules in adjescent monolayers [Erokhin, 1989]. This coordination is illustrated schematically in the Figure 6. Metal atom is in the center of the tetraeder formed by four oxygen atoms. Such coordination implies that the metal ions are bound to the fatty acid molecules in the adjacent layers and their attachment, very likely, takes place when the substrate pass through the meniscus during the upward motion. The other method of the monolayer transfer from the air/water interface onto solid substrates is illustrated in Figure 7. The method is called Langmuir-Schaefer (LS) technique (horizontal lift). It was developed in 1938 by I. Langmuir and V. Schaefer for the deposition of protein layers [Langmuir, 1938]. Specially prepared substrate touches horizontally the monolayer, and the layer transfer itself onto its surface. The method is often used for the deposition of rigid and protein monolayers. In both cases the application of LangmuirBlodgett method is not desirable as it results in the deposition of defective films.
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Figure 5. Scheme of the LB deposition: downward (a) and upward (b) motion of the solid support through the monolayer at the air/water interface.
Figure 6. Tetrahedral coordination of bivalent metal ions (central circle) by head-groups of adjacent monolayers.
In the case of the application of LS method to rigid monolayers special cares must be performed. The monolayer at the air/water interface must be divided into parts after reaching the desired surface pressure. It must be done with a special grid with windows, corresponding to the solid support sizes [Aktsipetrov, 1995]. The main reason of the utilization of the grid is the following one. If the monolayer is rigid, the removal of some its part will result in the formation of empty regions in the monolayer. As the layer is rigid, these empty zones will be maintained for a very long time. Repeating of the deposition will result in the formation of many defects in the monolayer, and the resulting transferred layer will be absolutely not homogeneous. The use of the grid provides also the guaranty, that only one monolayer is transferred during one touch. In the case of proteins (mainly, membrane proteins) the monolayer is soft [Erokhin, 2002]. Therefore, the problems, mentioned above, do not exist in this case and the use of the grid can be avoided. In fact, the monolayer structure in the case of protein layers is practically amorphous, what is easy to reveal by Brewster angle microscopy – a powerful tool for the monolayer domain structure visualization. Therefore, the removal of some monolayer regions can be rapidly compensated by the feedback system without the loss of the monolayer homogeneity. However, there is the other problem when applying the LS technique for the protein monolayer transfer. The situation on the solid support after the touching of the
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monolayer is schematically shown in the Figure 8. Regular closely packed monolayer is at the surface of the solid support. Some amount of water, transferred together with the monolayer, forms a drop at the substrate [Erokhin, 2000]. Some protein molecules can form not regular layer at the top of this drop. If the sample will be dried in a usual way, these molecules will form not homogeneous layer in not controllable way. Therefore, these additional molecules must be removed before the sample drying. The effective way to realize it is to use rather strong jet of inert gas, such as nitrogen or argon. It removes the water drop together with randomly distributed protein molecules on its top, leaving only regular layer, faced to the substrate surface.
Figure 7. Scheme of the LS deposition.
Figure 8. Scheme of the water and protein molecules distribution on the substrate surface after horizontal touching of the protein monolayer.
According to the transfer procedure, deposited films are usually divided into 3 types, schematically shown in the Figure 9, namely, X-, Y-, and Z-types [Roberts, 1990]. As it is clear from the Figure 9, the Y-type is a centrosymmetric one, while X- and Z-types are polar ones and differ one from the other by only the orientation of the head-groups and hydrocarbon chains with respect to the substrate surface. Such division appeared due to the fact, that in some cases there is no monolayer transfer during upward or downward motion of the substrate in the case of LB deposition. In the case of the LS deposition, moreover, the layers seem to be always transferred in a polar manner. However, the X- and Z-types are practically never realized in practice [Lvov, 1986]. Even if some nonlinear properties, such as
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pyroelectricity [Blinov, 1984], realizable only in polar structures, were observed, and the structures were considered as polar ones, detailed investigations revealed that the films are of Y-type with only not equal density of add and even layers [Roberts, 1989]. Therefore, thermodynamically stable structures, that in a case of LB films are bilayers, must be realized. Moreover, in the case of LS deposition of fatty acid films, it was shown that the last 3 transferred monolayers are involved into the structural reorganization during the meniscus passing, in order to realize thermodynamically stable Y-type packing [Kato, 1987]. This reorganization provides the orientation of the hydrocarbon chains to the air in the last deposited monolayer.
Figure 9. Types of the structures of LB films.
Figure 10. Fromherz trough. Circular trough is separated into sections (a). Monolayer formation and interactions with enzyme take place in different sections. The monolayer can be transferred from one section to the other by simultaneous motion of both barriers in the same direction due to the deformation of water subphase meniscus during passing the section walls.
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Generally, packing of amphiphiles in LB films is determined by the head-to-head and tail-to-tail packing. The tilting angle of the hydrophobic chains with respect to the film plane can be different, depending on the size of the head-group, but it must provide the close packing of the chains. However, several examples of the other packing with interdigitation of the hydrocarbon chains of adjescent layers were observed [Erokhin, 1989]. Significant increase of the interest to LB method appeared after the works published by the H. Kuhn group [Kuhn, 1970; 1972; 1979]. They had shown the possibility to realize molecular architecture with this method. Complicated structures with alternation of layers of different molecules were realized. Such systems were used for studying the energy transfer processes. Layers of donors and acceptors were separated by spacer layers of different thickness.
Lb Films of Enzymes: Fromherz Trough Application of the LB technique for the deposition of enzyme films meets the same difficulties as for the most of other proteins [Erokhin, 2002]. These difficulties are resulted from the organization of such objects. Practically all proteins (except membrane ones), including enzymes, have the most of charged groups at their surface, while hydrophobic groups are in their center maintaining the globular tertiary structure. Therefore, it is difficult to suppose that such molecule will remain in the native form at the air/water interface. Observed surface activity of protein molecules in the most of cases is mainly due to the complete or partial denaturation of the protein molecule. Hydrophobic interactions, maintaining the protein globule structure, are of the same order of magnitude as the action of the surface tension (72 mN/m for pure water). Thus, arriving to the air/water interface, protein molecules (including enzymes) must rearrange their structure. Hydrophobic groups will be oriented towards air, while charged groups will remain in water. Native protein globular structure is lost. Especially for enzyme molecules, such reorganization of the protein globule is very critical for the functioning. In fact, enzyme can be considered as an extremely effective catalyser and mutual orientation of specific groups, responsible for the improved electron transfer between involved reagents (substrates), is a key parameter. Even partial denaturation will result in the complete lost of the enzymatic activity. Above considerations have demonstrated the necessity of the modification of the traditional LB technique if we want to deposit enzyme-containing layers. Such modifications were developed basing mainly on the formation of complex layers, including lipid (or other surfactant) monolayers with attached enzyme layer. In this case the lipid layer is formed at the air/water interface. Its presence decrease the surface tension for the value of the reached surface pressure. Thus, enzyme molecules will be much less affected by the external forces when arriving to the air/water interface. However, in order to provide a stable complex between lipid monolayer and enzyme molecules, we need to involve some forces of interactions. The most frequently used interactions have the electrostatic nature [Peschke, 1987; Pachence, 1991; Edmiston, 1998]. Similarly to the case of LbL deposition, one need to determine the isoelectric point of the enzyme that is planed to be deposited. All the further manipulations will be performed at the pH, when the enzyme is charged and to find lipid
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molecule, capable to form stable monolayers and having an opposite charge in the headgroup at the chosen pH value. Thus, the procedure must be the following one. Lipid solution must be spread at the air/water interface and the monolayer must be compressed till the target surface pressure. After equilibrating, enzyme solution must be injected under the monolayer. It is important to inject the enzyme solution after the monolayer is already formed, otherwise enzyme molecules can be denatured during the contact with the pure water surface. Incubation time can vary from some minutes till some hours depending on the particular enzyme and lipid molecules as well as on the composition and pH of the subphase. The attachment of the enzyme molecules can be revealed by the variation of the surface pressure (if the feedback is switched off) or surface area, covered by the monolayer (if the feedback is switched on) as well as by measuring of the monolayer surface potential. After the incubation, such layer can be transferred onto solid supports using generally LangmuirShaefer technique (horizontal lift). A special device was designed for such purposes. It was developed by Fromherz and was called Fromherz trough [Fromherz, 1971; Fromherz, 1975]. The scheme of the device is shown in Figure 10. Usually, it has a circular geometry and is divided in several sections as it is shown in the figure 10a. Compression is provided by two barriers that are radii of the circular trough and move towards each other. Each section must have its own surface pressure sensor. Let us consider briefly the operation principles of this device. The formation of the lipid monolayer takes place in the section 1. The subphase can contain some additions (usually, salts), suitable to the formation of the stable layer. When the target pressure is reached and the monolayer is equilibrated, it must be transferred to the section 2 with the composition and pH facilitating the detachment of ions, attached during the previous stage, suitable for the stable monolayer formation but not useful for the interactions with the enzyme molecules. We can call this operation “two-dimensional washing”. The transfer of the monolayer from one section to the other is performed by the simultaneous motion of both barriers in the same direction. Then, the monolayer must be transferred to the third section, already containing enzyme molecules in the subphase volume. Adsorption of enzyme molecules onto the lipid head-groups takes place in this section. Composition and pH of the subphase must provide the most effective interactions. When the enzyme layer is formed, the complex layer will be transferred to the section 4 where the subphase is suitable to the layer transfer to the solid supports. In principle, it is possible to use not only electrostatic interactions, but some specific recognition [Uzgiris, 1987; Morgan, 1992]. In this case, the lipid molecules must have some groups providing high affinity to the other groups, already present or attached to the enzyme molecule surface. Of course, this approach is more complicated and expensive, as it demands synthesis stages. However, it can provide a desirable orientation of the enzyme molecules towards the analysed solution.
Special Modifications of LB Technique: Sliding Plate Method One other technical decision, suggested by V. Troitsky, allows to deposit enzymecontaining layers [Troitsky, 1996]. The method combines the LB deposition and self-
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assembling. Important feature of the method is a special construction of the sample-holder. Adsorption of the enzyme molecules can be performed on the lipid monolayer head-groups not only at the air/water interface, but also at solid supports. However, the substrate with deposited lipid layer must be transferred to the reaction medium (enzyme solution) without its exposition to air, otherwise not controllable reorientation of lipid molecules can take place. In fact, as it was demonstrated, last 3 monolayers of fatty acids reorient themselves by flip-flop transitions during the deposition by Langmuir-Shaefer technique (horizontal lift). Second important remark is connected to the necessity to avoid the contact of the enzyme molecules with air, what can result in the degradation of their properties. The developed construction of the sample holder allows to overcome these difficulties. The scheme of the sample-holder (sliding plate sample-holder) and principles of its functioning are shown in Figure 11. An essential part of the sample-holder is the presence of the sliding plate in the vicinity of the sample surface (about 1 mm). The plate is realized from hydrophilic material. It can be in two positions. In the “open” position the sample surface is exposed to the environmental medium. In the “close” position, water can be kept between the sample and the sliding plate by capillary forces. Figure 11 illustrates the deposition procedure realized by the described device [Pastorino, 2 002]. Sample with hydrophobic surface is attached to the sample-holder with sliding plate in “open” position. It pass through the already formed lipid monolayer at the air/water interface and the monolayer is transferred onto its surface with head-groups oriented to the water volume. In the bottom position, the sliding plate is transferred into the “close” state (Figure 11a). Thus, during the upward motion nothing will be transferred onto the sample surface and the water between the sample and the sliding plate will prevent the molecules in the monolayer from reorientation (Figure 11b). Then, the sample is transferred to the volume containing enzyme solution. The sliding plate is transferred to the “open” state and the adsorption of the enzymes takes place (Figure 11c). Similarly to the previous case, the enzyme-lipid interactions can have an electrostatic origin or can be based on high affinity of specific molecular groups. After the enzyme layer formation, the sliding plate is transferred to the “close” state and the sample is passed to the Langmuir trough. Still maintaining the plate in the “closed” state, the sample must go through the lipid monolayer till the bottom position. In this position, the sliding plate is transferred into the “open” state and the sample passes through the lipid monolayer during upward motion. Transferred lipid layer provide the protection of the enzyme molecules from the exposition to the air (Figure 11d). Figure 11e demonstrates the motion of the sliding plate sample-holder between different sections of the equipment. The effectiveness of the method was initially demonstrated for the layers containing cytochrome P450 [Troitsky, 1996]. Then, using electrostatic interactions the method was successfully applied for the deposition of enzyme-containing films, in particular, films containing penicillin G acylase [Pastorino, 2002]. It is to mention, that the temporal stability of enzymes in such protected layers was shown to be significantly higher with respect to the enzymes in solution and in films prepared by traditional immobilization techniques.
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Figure 11. Deposition method using sliding plate sample-holder. Figures 11a-11d represent different stages of the enzyme-containing structure formation, while Figure 11e shows the motion of the sample-holder between different sections of the deposition equipment.
Langmuir Monolayers for Study of Enzyme-Membrane Interactions Monolayer at the air/water interface can be considered as a half of a model biological membrane. Its composition can be varied allowing to model different situations. Moreover, variation of the surface pressure can allow to identify different membrane regions that are more affected to the enzyme activity actions. Different measurements can be performed during such investigations. Practically in all cases when the interaction of enzymes with Langmuir monolayers is considered, measurements of the variation of the surface pressure and/or monolayer area are carried out. Usually, the monolayer of a desirable composition (containing natural or synthetic lipids) is spread at the air/water interface and compressed till the target pressure. Then enzyme solution must be added to the water subphase. Measurements can be performed when the feedback, maintaining the fixed surface pressure, is switched on or off. In the case of the switched on feedback, one will observe the variation of the monolayer area during the interaction. Two distinct situations of the enzyme effect on the monolayer can occur. They are schematically illustrated by the Figure 12.
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Initial organization of the lipid monolayer at the air/water interface is shown in Figure 12a. In the case of the increase of the monolayer area (Figure 12b), one can conclude that the enzyme is attached to the monolayer and, in some cases, can even partially penetrate into it. Instead, it is possible to observe also the decrease of the monolayer area. It can mean that the enzyme action on the model membrane result in the formation of local collapses and/or lipid micelles, that then leave the air/water interface and penetrate the aqueous subphase. This situation is illustrated by the Figure 12c. Of course, more precise interpretation of what really happened will demand additional investigations with other, more powerful techniques. Some of such useful technique will be mentioned below. However, even the information on the monolayer area variation can be very useful and can allow even quantitative estimation of the process. In the case of the switched off feedback, the monolayer area is fixed and one can register the surface pressure variation during the enzyme-model membrane interactions. In principle, the information obtained during such investigations is comparable with that in the case of switched on feedback. However, the difference is that in the case of “feedback on” situation the state of monolayer is maintained constant during the interactions, while in the “feedback off” case the variation of the surface pressure can result in the phase transitions of the monolayers state (for example, from liquid expanded to liquid condensed). Therefore, the interactions of enzyme molecules with the monolayer can be varied from the beginning till the end of the process. The other possibility to study these interactions is to measure the surface potential variations. As it was mentioned above, the measurements of the surface potential is usually performed with the Kelving probe and can provide the information of the charges and dipoles distribution across the monolayer. Therefore, in some cases the variation of the surface potential can indicate the attachment of the enzyme to the monolayer (as it contains its own charges and dipoles) as well as reorganization and reorientation of the molecules in the monolayer. In this last case the conformational changes of the lipid head-groups will be more evident as their polar nature result in a significant contribution to the total value of the surface potential. Considering the morphology of the monolayer and its variation during the interactions with enzyme molecules in the subphase, two types of microscopies are widely utilized. Fluorescence microscopy [Lösche, 1984; Chi, 1987; Schwartz, 1993] is based on the addition of the fluorescently labelled probes to the system under investigation. The system in this case must be illuminated with the light of the wavelength, corresponding to the fluorescence excitation. Imaging must be performed after filtering of the exciting light. These dye molecules can be added to the spreading solution of lipids. During the monolayer compression, the dye molecules will not be able to penetrate regions with lipid molecules close packing, visualizing therefore the domain structure of the model membrane. The domain sizes and shape can be varied during the interaction with enzymes and these morphology variations can be easily visualized even in a real time. The other possibility is to work with a monolayer without dye addition, but to use fluorescently labelled enzymes. In this case it will be possible to determine and to localize the interaction of the enzymes with the monolayer registering the appearance of the fluorescence.
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Figure 12. Lipid monolayer at the air/water interface (a). The interaction with enzyme molecules in the water subphase can result in the protein adsorption at the lipid head-groups and even in their partial penetration into the monolayer and, therefore, in the increase of the monolayer area at the given value of the surface pressure, maintained by the feedback system (b). In other cases, interactions with enzymes can result in the local monolayer collapses and partial transfer of lipid molecules into the volume of the water subphase in the form of micelles (c).
Second type of the microscopy, widely used for the monolayer investigations, is called Brewster angle microscopy (BAM) [Overbeck, 1994; Tsao, 1995; Angelova, 1995]. It is based on the fact that polarized light, incident to the interface at the Brewster angle, is not reflected. Thus, BAM instrument is equipped by goniometric system with attached laser and CCD device, allowing to vary angle of incidence and, accordingly, reflectance. Usually, the incidence angle corresponds to the Brewster angle for the air/water interface. Thus, the pure water surface will be seen as the dark field in the acquired image. The presence of the monolayer will vary Brewster conditions for both water/monolayer and monolayer/air interfaces resulting, therefore, in the appearance of bright areas in the image, corresponding to the position of monolayer domains. It is clear that this technique can be also used for the study of interaction of enzymes with model membrane, as it will allow to visualize the variation of the domain sizes and shape during such interactions. In addition, BAM has one significant advantage with respect to the fluorescence microscopy. In the case of BAM we do
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not need to add any dye or other molecules into the system under investigation. In other words, the system in the case of BAM is much less affected by the artefacts added for the visualization possibility in the case of fluorescence microscopy. Other technique that is used for the monolayer study and can be useful for the investigation of the interaction of enzymes with model membranes is ellipsometry [Feachem, 1934; den Engelsen, 1974; Ducharme, 1985; Kim, 1990]. The variation of the polarization state of light reflected from the interface can give the information about the variation of thickness and refractive index of the thin layer at the interface. However, it seems even more effective to use X-ray reflectivity measurements for these reasons [Helm, 1987; Kjaer, 1988; Majewski, 1998; Kaganer, 1999]. Modelling of the experimental curves can give the information not only about the thickness of the layer, but even to reconstruct the electron density profile in the direction normal to the monolayer plane. In fact, in the case of the monolayer interaction with DNA it has allowed to register not only the fact of the DNA adsorption, but also the state of DNA in complexes with different lipids [Erokhina, 2007; Cristofolini, 2007]. In the case of interactions with enzymes, such measurements can help in more precise localization of the enzymes in model membranes during the interaction processes. Effective utilization of the X-ray reflectivity measurements demand the use of synchrotron radiation as it provides high signal-to-noise ratio, necessary for the reliable model construction. As the most serious drawback of the method, we can mention that it is rather time consuming and does not allow real time monitoring of the process. However, current works in the field, especially based on the analysis of the diffuse scattering allow to forecast that in the nearest future these limitation will be successfully overcame. Of course, as more techniques are available for the characterization, as better and precisely the interaction process will be described. Just to illustrate the previous considerations, let us consider some specific example of the application of the monolayer technique for the investigation of the interactions of enzymes with model membranes. Historically, these examples will correspond to the first peak of the activity, shown in Figure 1, when the effectiveness of such studies was well demonstrated. The most of the current studies are based on the same approaches and assumptions, as it was 30 years ago, with the only difference that much more powerful characterization techniques are available. Investigation of the hydrolysis of the lecithin monolayer by Crotalus adamanteus αphospholipase A2 had allowed to demonstrate that the active dimeric form of the enzyme is in equilibrium with inactive subunit. The study had also demonstrated that the calcium ions presence is required for the surface reaction. Thus, authors have demonstrated the applicability of the monolayer technique for studying lipid-enzyme interactions [Lagocki, 1970]. In the other study similar system, containing lecithin monolayer at the air/water interface and phospholipase A in the subphase, was investigated in order to identify the relative position and orientation of the hydrophilic groups of the various membrane constituents [Colacicco, 1971]. These work has two essential features. First, it was explicitly shown that the measurement of the surface potential can provide a very important information about the interaction of lipid molecules in the monolayer with enzymes in the water subphase. Second,
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it was demonstrated that the kinetics of the interaction processes depends on the initial surface pressure of the monolayer and, therefore, on the state of the model membrane.
Enzyme-Containing LB Films as Sensitive Layers of Biosensors We have attributed the second increase of the activities in the field of enzyme-containing LB films (Figure 1) to the possibility of using these layers for molecular and biomolecular electronics and as sensitive elements of enzymatic biosensors. Several specific features of the deposition method have attracted the increased attention as practically no other method can provide similar possibilities. First of all, in the case of the LB technique we can form a film, containing only one molecular layer of the enzyme. This feature can be very important for the industrial applications when we have to work with expensive enzymes. Second, protection of the enzyme sublayer by its incorporation into the lipid layers can, as we have discussed above, increase significantly its temporal stability, preventing the contact with undesirable surrounding containing polutants. Third, the LB method allows to realize complex structures where enzyme layers can be included into the structures, containing also layers of conducting molecules that can be considered as mediators, facilitating electron transfer between the enzymes, substrates and products. In the most of cases, conducting polymers, such as polyaniline, polythiophene, etc., are used for these purposes. Let us consider particular examples of the utilization of the enzyme-containing LB films as the sensitive layer of biosensors. Glucose oxidase (GOD) is probably the mostly used enzyme as its practical usefulness is obvious [Okawa, 1989; Rosilio, 1997; Anicet, 1998]. The working principle of the biosensors for the glucose detection is based on the following reaction: GOD GLUCOSE
GLUCONIC ACID + H2O2
Thus, sensitive GOD layer must be deposited onto the working electrode. The sensor chamber must contain also reference electrode, that is usually realized from Ag-AgCl material. It must also contain counter electrode, usually realized from Pt, that must provide electrical carriers necessary for the maintaining the potential difference between reference and working electrodes and not to be involved directly to the electrochemical reactions. Therefore, the current in the measuring circuit will be proportional to the concentration of the hydrogen peroxide and, therefore, to the concentration of the glucose in the solution under the investigation. We must also consider several possibilities that can improve the performance of the sensitive enzymatic layer and, therefore, the performance of entire biosensor. First of all, the GOD layer is deposited always together with the surfactant monolayer. For some particular reasons, easiness of the glucose access, for example, the presence of this surfactant layer can be undesirable. It was shown the possibility to remove such layer after the active layer formation practically without the damage of the enzyme properties in the active sensitive layer. In particular, when the GOD layer was transferred together with
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behenic acid monolayer, the removal of the last was possible by isopropanol treatment of the complex layer. It is to note, that such selective removal of one compound of the formed complex layers is rather well-known and was used for the removal of fatty acid molecules from the layers leaving only fatty acid salt molecules at the solid support surface. This process was called skeletonization process. It is very interesting itself, as it allows to form the layer with desirable porosity. In fact, the ratio of the fatty acid – fatty acid salt molecules in the layer can be easy controlled by the pH variation. After the layer deposition, this ratio is preserved in the film. Organic solvent treatment will remove only soluble counterpart (acid) maintaining, however, the overall structure of the film. Therefore, desirable fraction of cavities will be realized. Other example of the selective removal of one compound from the complex layer is connected to the formation of aggregated inorganic nanoparticulate layers starting from LB organic precursors. As it was shown, the method allows to remove organic molecules selectively from the layer, leaving on the support surface only ultrathin inorganic films [Facci, 1994; Erokhina, 2002]. The accuracy of the film thickness realization in this case can reach the value of about 0.5 nm what is comparable with the most developed, complicated and expansive techniques, available now, such as molecular beam epitaxy. As it was mentioned above, the method allows also to deposit complex layers, where GOD can be attached to conducting polymer sublayers instead of traditional lipids or surfactants. Such complex layers were shown to exhibit the improved properties when they are used as sensitive layers. Finally, let us present an overview of the enzymes involved into investigations using Langmuir and Langmuir-Blodgett films. About 200 works were published where different lipases were involved into the study. The fact does not seem strange if we consider the membrane-active properties of these enzymes. Here we present references to only some representative works with these enzymes (Lagocki, 1970; Colacicco, 1971; Shen, 1975; Bianco, 1989; Davies, 1991; Ivanova, 1993; Mirsky, 1994; Grandbois, 1999; Estrela-Lopis, 2001; Jensen, 2002; Nielsen, 2002; Pastorino, 2002; Wang, 2005). Glucose oxidase is the other enzyme widely studied applying Langmuir technique. About 50 articles were published on this object (Okahata, 1989; Sun, 1991; Lee, 1993; Zaitsev, 1993; Rinuy, 1999; Zhang, 2000; Zayats, 2002; Watanabe, 2005; Lee, 2007). Rather low cost and obvious usefulness of this enzyme for sensor applications are main reasons for such activities. Other relatively widely studied enzymes are peroxidases (Chasovnikova, 1992; Razumas, 1996; Berzina, 1998; Tang, 2002; Morandat, 2004), urease (Langmuir, 1938; Gidalevitz, 1999; Hou, 2002; Singhal, 2002) and acetylcholinesterase (Choi, 1997; Dziri, 1999; Choi, 2001; Wang, 2006). Other enzymes to whome Langmuir technique was applied are: ATPase (Babakov, 1979; Prokop, 1995; Wu, 2001), luciferase (Marron-Brignone, 1996; Marron-Brignone, 2000; Palomba, 2006), cytochrome c oxidase (Tredgold, 1979; Salamon, 1993; Cullison, 1994), glutamate dehydrogenase (Hourdou, 1995; Girard-Egrot, 1997), hydrogenase (Nakamura, 1998; Noda, 1998; Qian, 2002), catalase (Nitsch, 1990; Maksymiw, 1991), cholesterol oxidase (Slotte, 1992; Slotte, 1993), choline oxidase (Girard-Egrot, 1997; Girard-Egrot, 1998), cholinesterase (Danilov, 1975; Godoy, 2005), monoamine oxidase (Barmin, 1994),
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tyrosine hydroxylas (Eremenko, 1991), protease (Gole, 2000; Vinod, 2007), alkaline phosphatase (Petrigliano, 1996; Giocondi, 2007), alcohol dehydrogenase (Pal, 1994), alphachymotripsin (Anzai, 1989), anhydrolase (Mello, 2001, Mello, 2003), cutinase (Ivanova, 2003), DNA polymerase (Mizushina, 2000; Matsuno, 2001), RNA polymerase (Rajdev, 2007), DNA gyrase (Lebeau, 1990; Lebeau, 1992), glutathione S-transferase (Paddeu, 1996), penicillin G acylase (Pastorino, 2002), sphingomyelinase (Harte, 2005; Rao, 2005), xylanase (George, 2002), endonuclease (Matsuo, 2005), glycosyltransferase (Nagahori, 2003), hexokinase (Castro, 2007), human erythrocyte catalase (Harris, 1995), malate dehydrogenase (Peters, 1975), nuclease (Inbar, 1976), phytase (Caseli, 2006), saccharase (Sobotka, 1941), transglutaminase (Faergemand, 1997), trypsin (Fromhertz, 1975), xanthine oxidase (Kristensen, 1998).
The Electrostatic Layer by Layer Self-assembly Technique General Principles The LbL technique was firstly introduced by Iler in 1966 for the alternate assembly of oppositely charged layers of colloidal particles, such as silica and alumina [Iler, 1966]. In the 1990’s Decher and co-workers reported the fabrication of multicomposite films of charged materials trough LbL adsorption from aqueous solutions [Decher, 1991; Knoll, 1996; Decher, 1997]. Since then, extensive work has been carried out on the application of this technique to the fabrication of multilayered ultrathin films incorporating a broad range of charged macromolecules including synthetic polyions, biopolymers, viruses, ceramics and nanoparticles [Lvov, 1995; Ariga, 1997; Caruso, 1997; Lvov, 2000; Salditt, 2002].
Figure 13. Scheme of the LbL deposition procedure, based on the successive adsorption of polyanion and polycation layers on the solid supports.
The general principle of the LbL technique is very simple and, as already mentioned, is based on the alternate adsorption of oppositely charged species on the surface of a charged solid support. Figure 13 schematically describes the assembly procedure.
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In the figure a negatively charged support is dipped into a diluted aqueous solution/dispersion of a cationic specie for a fixed period of time, which has to be experimentally determined in order to reverse the charge of the support surface. As a result, a thin layer of the cationic specie is formed on the support, thereby generating a positively charged surface. The positively charged support is then rinsed in water, in order to remove the unbound material, and dipped into a diluted aqueous solution/dispersion of an anionic specie for a given period of time. The rinsing step is repeated and the support, which is now negatively charged can be used again for the assembly of a successive positively charged layer. By repeating the described steps, a stable and uniform multilayered structure can be deposited with a desired thickness ranging from few nanometers to 10 μm, with precision better than 1 nm and with a predetermined structure and function. The driving force of the assembly process is the electrostatic one, but other interactions can be utilized such as covalent bonding [Schultz, 2005; Bai 2006; Lu, 2007] and biomolecular recognition [Sano, 1996; Cassier, 1998; Dai, 2007;]. The physico-chemical properties of the multilayer architectures can be modified by varying the deposition conditions, such as pH and ionic strength of the solutions [Voigt, 2003; Schonhoff, 2003]. Finally different charged species can be utilized in the same assembly for the fabrication of complex structures. What makes this technique so promising is that it does not require the use of complex equipments, as the assembly in the laboratory is carried out in beakers, the use of harmful regents, as the assembly is carried out from aqueous solution and finally the assembly is not limited to flat surfaces but it can be carried out onto 3D supports of any shape. The above considerations marks clearly the LbL technique as a powerful tool also for processes to be conducted at the industrial scale considering that the simplicity of the technique implies that no complex industrial plant would be required, that this technique can be regarded as environmentally clear and that its versatility guarantees applications in a wide range of technological fields.
LbL Films of Enzymes: Structure and Properties Assembly Procedure on Planar Supports Multilayers containing biomolecules, including enzymes have been fabricated by means of the LbL technique and their functionality has been characterized [Lvov, 2000; Lvov, 2002; Lvov, 2003]. The first publication on enzyme LbL multilayer formation was published by Lvov et al in 1995 [Lvov, 1995] and assembly of cytochrome, myoglobin, glucose oxidase and peroxidase in alternation with oppositely charged linear polyelectrolytes and nanoclay was described. Proteins in the LbL multilayer preserved their conformation and enzymatic activity. Regular multilayers of different proteins were also prepared forming by this protein supperlattices. The starting point in the set up of an assembly protocol for the use of each particular protein, is represented by the determination of its isoelectric point. The working pH of all the solutions has to be set apart from this value so that the protein molecules are sufficiently positively or negatively charged. It is interesting to note how the same protein can be utilized
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either as a positively or negatively charged specie [Liang, 2005]. When a decision has been made on the charge to be adopted for the protein molecules, the polyions to be used in the assembly process can be defined and thus the architecture of the multilayered structure. For a successful assembly of proteins, it is important to alternate the protein layers with linear or branched polyions as it has been demonstrated that flexible polyions penetrate the protein layer acting as an “electrostatic glue” [Lvov, 2000]. The following steps can be individuated in the fabrication of a protein containing multilayer on the surface of a planar support: (1) prepare the protein and polyions solutions at a concentration between 0.1 and 1 mg/ml and between 1 and 2 mg/ml respectively at the established pH (1 or 2 units below or above the isoelectric point of the protein under consideration). When working with expensive proteins it is important to use as less material as possible in a single experiment, in this case the protein solution concentration can be kept at lower values and the adsorption time consequently has to be longer; (2) prior to start the protein multilayer fabrication, take your planar support having a charged surface (e.g. silicon, quartz, glass slides) and deposit on it minimum three polyion layers in order to provide an uniform surface with a well defined charge. It has been demonstrated that precursor films are necessary to provide liner mass increase for the following layers [Lvov, 1995]. Let’s consider as an example the case of a negatively charged support. During the first step it is dipped into a water solution of a polycation, such as poly(dimethyldiallylammonium chloride) (PDDA), at a concentration of 2 mg/ml for 10 min and then it is rinsed for 1 minute using a buffer solution at the same pH of the solutions used in the assembly in order to keep the polyions and protein ionised. The same procedure is then repeated for a polyanion solution, such as poly(styrenesulfonate) (PSS). Repeat the alternate adsorption of PDDA and PSS in order to deposit a two bilayer precursor. For the subsequent protein adsorption, the precursor will have PDDA or PSS as outmost layer depending on the charge of the protein. (3) the modified support bearing the right charge for protein adsorption is now dipped in the protein solution for a time between 10-60 min depending on the protein concentration [Caruso, 1997; Brynda, 2005; Lu, 2007]. As proteins generally are not stable at ambient temperature, it is recommendable to carry out the protein adsorption step at 4 °C. After rising the sample, (4) it is dipped in the solution of the oppositely charged polyion for further deposition. The steps can be repeated for the deposition of the desired multilayered structure. The final structure is then stored in buffer solution preferably at 4 °C. This is a general procedure which applies to the fabrication of protein containing multilayers and which has to be experimentally tailored to the specific molecules and biomolecules under the use.
Structure Characterization of LbL Films on Planar Supports The progressive build-up of the multilayered films onto a planar support can be monitored using different well-established techniques such as UV-Vis spectroscopy, quartz crystal microbalance (QCM) and surface plasmon resonance (SPR). The easiest way to characterize the deposition process is represented by the employment of UV-Vis spectroscopy. In this case, UV-Vis absorption spectra of the film deposited onto a
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quartz slide are recorded after each layer or bilayer deposition. The multilayer growth is then monitored by the enhancement of the absorbance peak intensities of the deposited species. Typically proteins display an absorbance peak around 280 nm, a linear increase of the adsorbance at this wavelength, plotted against the number of deposited layers, would confirm a proper and uniform deposition. Another useful technique, which does not require expensive and complicated instruments, is QCM. QCM is based on the properties of piezoelectric quartz crystals to vary frequency of their resonant oscillations according to the state of the resonator surface. The deposition/adsorption of molecules onto the surface of the quartz determines a decrease in the oscillating frequency. A direct relation between the frequency shift and the deposited mass can be obtained using the Sauerbrey equation [Sauerbrey, 1959; Facci, 1993]: Δf = - Δm x C where C Δf f0 Δm A ρq μq
= 2f02 / A(ρqμq)½ = measured frequency shift, = resonant frequency of the fundamental mode of the crystal, = mass change per unit area (g/cm2), = piezo-electrically active area, = density of quartz = shear modulus of quartz
The Sauerbrey equation can also be expressed as: ΔL = -Δf / Cρ where ρ = density of the deposited material. The density of the protein / polyion film has been found to be approximately 1.3 g/cm3 [Lvov, 1995]. QCM is a very sensitive tool to detect changes in mass and thus a helpful method to sense adsorption processes both at solid/gas or solid/liquid interfaces. For measurements at solid/gas interface, the assembly process is carried out by dipping the quartz crystal in the adsorption solution for the given amount of time then, after the rising step, the quartz crystal is dried in a nitrogen stream and the resonance frequency shift is measured. For measurements at the solid/liquid interface, only one electrode is usually used. In this case the quartz crystal is mounted in a flow chamber where one electrode is placed in a permanent contact with the adsorption solutions while the electrode on the other side is maintained in the air. In the case of the necessity of the utilization of both oscillator electrodes, each of them must be placed in separate reaction chambers, avoiding possible
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short circuit through the solution that can result in the significant decrease of the quality factor and, therefore, the accuracy of the measurements. A regular stepwise increase in frequency shift, and consequently in film mass and thickness, with the number of assembly steps indicates a successful deposition procedure. The QCM technique is a standard technique usually used in order to elaborate the assembly procedure, by defining the time required to reach the saturation adsorption of the deposited species. SPR can be also used to investigate the construction of LbL films and to quantitatively monitor the assembly process. SPR is based on the property of the significant transfer of the incident light energy to the oscillations of the free electron plasma (surface plasmons) in thin metal layers. The resonance angle (minimum of the reflected light) is strongly dependent on the state of the metal/air or metal/liquid interface allowing to consider the technique as very useful one for the deposition process monitoring. Depending on the thickness of a molecular layer at the metal surface, the SPR phenomenon results in a graded shift of the angle corresponding to the significant reduction reduction in intensity of the reflected light [Frutos, 1998]. To clarify the structure and morphology of the LbL assemblies, the multilayer films are generally examined by atomic force microscopy (AFM) and scanning electron microscopy. The atomic and molecular composition of the assemblies can be determined using X-ray photoelectron spectroscopy (XPS) and Fourier transform infrared spectroscopy (FTIR).
Properties of Lbl Films Containing Enzymes Films of biomolecules assembled with strong polyions have been found to be insoluble in buffer solutions for pH values from 3.00 to 10 [Lvov, 2002]. Bioactive molecules such as enzymes, included into the structure, are not denatured by the deposition process demonstrating thus the potentiality of the LbL technique in the area of biocatalysis [Caruso, 2000a; Yu, 2005; Patel, 2005; Rodriguez, 2006; Hamlin, 2007; Szabo´, 2007]. The versatility of the technique allows the fabrication of biocatalysts with a tailored catalytic activity. This can be achieved by varying the number of enzyme layers, their position and sequence in the multilayer [Pastorino; 2003]. Moreover, it has been demonstrated that in some cases enzymes embedded in such multilayers are more stable than free enzymes. The explanation for this stability improvement can be attributed to the ordered arrangement of the enzyme molecules which protects them from microbial attack and from changes in microenvironment [Onda, 1999]. As relates to the residual catalytic activity for the LbL immobilized enzymes, it has been found to vary from 20% up to 60% of the catalytic activity of the free enzyme, depending on the enzyme under study [Pastorino, 2003; Stein, 2003; Linag, 2005]. The observed decrease in the catalytic activity is mainly due to substrate diffusion limitation in the multilayer and to the difficulty in reaching the active site of the immobilized enzyme molecules. The influence of diffusion limitation can be limited by increasing the ionic strength of the adsorption solutions or of the catalytic assay solution. At low ionic strength, polyions are strongly charged forming thus compact deposited layers, whereas at high ionic strength the polyion charges are partially neutralized resulting in a coiling conformation of the deposited layer [Lvov, 1994]. Consequently the substrate molecule can more easily
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penetrate the multilayer and reach the enzyme active site. Linag at al. [Liang, 2005] demonstrated that urease immobilized by LbL technique increases its catalytic activity from 23% to 65% (with respect to the catalytic activity of the free enzyme) as a consequence of the addition of 0.05 M NaCl in the assay solution. Another way to limit the influence of substrate diffusion is represented by the use of high surface area supports for the multilayer deposition [Onda, 1996a; Onda, 1996b]. As already outlined, the LbL technique is not limited to planar surfaces but can be used for the assembly on 3D supports. Yu et al [Yu, 2005] performed the LbL deposition of peroxidase-PSS complexes and oppositely charged poly(allylamine hydrochloride) onto porous polycarbonate membrane (pore diameters 400 or 100 nm) for the preparation of high surface area biocatalytic thin films. Activity enhancements of up to almost an order of magnitude larger were observed for the enzyme films deposited on the membranes compared with identical films formed on planar supports with the same geometrical area. The possibility of employing cellulose microfibers as a support to fabricate bioactive composites with organized enzyme multilayers was also demonstrated [Xing, 2007]. Through the LbL nanoassembly, laccase and urease were sequentially deposited with polycations on cellulose microfibers. The catalytic activity of the biocomposites was found to be about 50% of the initial activity after 2 weeks for laccase and after one week for urease (storage temperature at 4 °C). The catalytic activity was proportional to the number of coated enzyme layers. Such enzyme modified biocomposites could be used to decompose urea or lignin, or synthesize inorganic nanoparticles or polyphenols. Moreover urease fiber biocomposites were successfully applied for biomineralization to grow calcium carbonate microparticles needed for paper whitening. Li et al. [Li, 2006] have proposed the preparation of nanoporous polyelectrolyte multilayer films to be used as nanomembranes for the immobilization of proteins. Nanoparticle-polyelectrolyte hybrid multilayers were assembled by the alternate adsorption of anionic blend of silica nanoparticles and poly(acrylic acid) (PAA) and a cationic polyelectrolyte, poly(allyamine hydrochloride) (PAH). After chemically cross-linking the PAA/PAH within the films, the nanoparticles were removed by dissolution with hydrofluoric acid and a porous polymer film was obtained. The adsorption of bovine serum albumin onto the porous film was characterized and demonstrated to increase with increasing number of bilayers. Pollutants, impurities, interfering substances, or aggressive media can result in a lack of desirable properties and system integrity and may cause serious problems in the usage of such biocatalytic systems. Oxidation is one of the serious problems for the devices based on substances originated from living organelles as reactive oxygen has the ability to damage biomolecules. The oxidation leads to inhibition of their activity and changing of their properties. Shutava et al. developed an approach to protect substances encased in LbL films [Shutava, 2006]. This approach is based on the deposition of a layer which is permeable for the molecules of interest and acts as a barrier for pollutants on the top of the biocatalytic film of interest. Namely, hemoglobin/polyelectrolyte films were adopted as model systems, as hemoglobin has some biocatalytical properties and can be irreversibly destroyed by hydrogen peroxide. It was demonstrated that a catalase layer deposited on the top of the multilayer acts as a protective antioxidant barrier lowering the amount of hydrogen peroxide reaching the
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hemoglobin layers in the film depth. Hemoglobin in such films retained its nativity for a longer period of time. This was found to be a successful strategy for the protection of LbL assembled films. The LbL technique provides opportunities to tune the protein content in nanostructured films, which is of pivotal interest for applications in biocatalysis. It was shown to be possible to incorporate enzymes by LbL technique into the cellulose fiber structures, that cab be considered also as nanostructured films [Xing, 2007]. Other important applications of enzyme-containing LbL layer are in the field of sensor development, where these layers can be used as sensitive element for the detection of different compounds [Shutava, 2004; Karnati, 2007].
Nanobioreactors Two different approaches have been developed for the fabrication of nanobioreators by the LbL technique. The first approach is based on the assembly of enzyme multilayers on mico- and nano- colloidal particles (Figure 14). Enzyme multilayers on a particle shell have been found to be attractive for biocatalytic applications due to their potential to yield higher enzymatic reaction efficiencies than their planar film counterparts. This approach, for the immobilization of proteins, was firstly introduced by Caruso et al [Caruso, 1999] who demonstrated the ordered assembly of shells containing such proteins as bovine serum albumin and immunoglobulin G onto the surface of 640 nm diameter polystyrene latex particles. Since then different proteins, including enzymes, have been immobilized following this approach. Multilayered nanoreactors with β-glucosidase [Caruso, 2000a], glucose oxidase [Schuler, 2000; Fang, 2002], horseradish peroxidase [Caruso, 2000b], manganese peroxidase [Patel, 2005], lignin peroxidase [Patel, 2005], lactate oxidase [Stein, 2003], luciferase [Pastorino, 2003], urease [Lvov, 2001a; Lvov, 2001b; Liang, 2005] have been fabricated and characterized, demonstrating also directly electrochemical reactions [Lvov, 1998; Kong, 1998].
Figure 14. Colloidal particle covered by LbL layer with incorporated enzyme molecules.
Let us consider briefly the procedure for the multilayer shell formation. A known number of particles are added to a centrifuge tube followed by the sequential addition of polyions and
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enzyme solutions to fabricate the shell of required architectures. After addition of the polyions/enzyme an experimentally determined time is allowed to elapse so that saturation adsorption of the polyions or enzyme on the colloid particles is reached. The coated latex spheres are then centrifuged and the supernatant containing the unadsorbed species is removed. The washing procedure, including the adding of the pure water to the precipitate, shaking of the solution for several minutes, centrifugation and water removal, is generally repeated three times after the adsorption step to avoid admixing of the sequentially deposited components. Shell formation on colloidal particles can also be achieved by surface controlled precipitation [Dudnik, 2001]. Heterocoagulation process of polymer at the suspension of colloidal particles is a way for the controlled coating of particles. Precipitation can be achieved by non-soluble complex formation between polyelectrolyte and multivalent ions or by mixing the polymer solution with non-solvent [Radtchenko, 2002].
Figure 15. Encapsulation by LbL covering of the enzyme microcrystal or microaggregate template with polyelectrolyte layers.
Due to the versatility of the technique, it is possible to include into the shell also nanoparticle layers to provide enhanced functionality [Caruso, 2000b; Lvov 2001a]. Organized multilayers of nanoparticles and glucose oxidase were deposited in alternation with oppositely charged polyelectrolytes on 420 nm latex particles [Fang, 2002]. The inclusion of silica nanoparticles resulted in the higher glucose oxidase adsorption and thereby increased catalytic activity. The presence of magnetic nanoparticles allowed self stirring of the nanoreactors with a rotating magnetic field. Moreover, the magnetic properties of the nanoreactors can be used to efficiently separate them from the reaction medium [Caruso, 2000b] in the perspective of an industrial application of such biocatalysts. The second approach, which can be adopted for the fabrication of nanobioreators by means of the LbL technique, is based on the encapsulation of enzyme molecules into nanoshells (Figure 15). This can be achieved by the deposition of polyions onto enzyme microcrystal or microaggregates templates. The encapsulation of the enzyme catalase crystals into a multilayered shell [Caruso, 2000c; Jin, 2001] was reported as the first example of biocrystal templated assembly of polyelectrolytes. The method took advantage of the fact that catalase
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is a crystalline suspension in water at pH 5-6 and therefore can be treated as a colloidal particle. Catalase crystals (8x12 μm), which exhibit a positive surface charge at pH 5, were used as templates for the sequential deposition of PAH and PSS. By exposing the encapsulated enzyme crystals to a solution of pH 2, the crystals were solubilized, which occurs because the polyelectrolyte multilayers are permeable to small molecules in solution. The capsules were not destroyed after crystal solubilization indicating their high stability [Caruso, 2000c]. Multilayer films of polyelectrolyte-encapsulated catalase microcrystals, multilayer films of polyelectrolyte-uncoated catalase microcrystals and multilayer films of polyelectrolyte-solubilized catalase were prepared and their catalytic activity was compared. Prior to the measurement of the catalytic activity, the films containing both coated and uncoated crystals were immersed in a buffer solution at pH 7.0 for 2 h to solubilize the enzyme crystals ( catalase can be solubilized at pH< 4 and pH> 7). Multilayer films of polyelectrolyte-encapsulated catalase exhibited a catalytic activity 10 and 50 times higher than those of the corresponding films of polyelectrolyte-uncoated catalase microcrystals and multilayer films of polyelectrolyte-solubilized catalase [Jin, 2001]. The preparation of protein-loaded CaCO3 microparticles and their use as deposition templates was also reported as a way to encapsulate proteins [Antipov, 2003] (Figure 16).
Figure 16. Loading of porous microparticles with enzymes with their successive encapsulation by polyelectrolyte layers and template removal (dissolving of CaCO3 microparticles).
Figure 17. Loading of hollow polymeric capsules with enzymes by pH-driven opening and closing of pores in the capsule shell.
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Physical adsorption of proteins from the solutions onto preformed CaCO3 microparticles, and protein capture by CaCO3 microparticles in the process of their formation were proposed. The latter was found to be about five times more effective than the former (~100 vs ~20 μ g of captured protein per 1 mg of CaCO3). The enzymatic activity of α-chymotrypsin captured initially by CaCO3 particles during their growth and then recovered after particle dissolution was found to be about 85% compared to the native enzyme. Core decomposition and removal after assembly of the required number of polyelectrolyte layers resulted in release of protein into the interior of polyelectrolyte microcapsules [Petrov 2005]. Another way to achieve the encapsulation of enzyme molecules is represented by the possibility to obtain hollow nanocapsules by the LbL technique as demonstrated by Donath et al [Donath, 1998; Caruso, 1998] and by loading them with enzyme molecules (Figure 17). This approach is based on the formation of a multilayer film onto the surface of a template with its subsequent dissolving by the variation of pH or composition of the environmental solution [Mohwald, 2000; Sukhorukov, 2001; Antipov, 2004]. Different templates can be used to this aim such as organic, inorganic cores and biological cells. The template decomposition is achieved by different means depending on the template. Carbonate particles, such as CaCO3 and MnCO3 particles can be easily dissolved in 0.01 M HCl [Shchuki, 2004], melamine formaldehyde can be dissolved in 0.1 M NaCl and in solvents such as DMSO [Hai, 2004], biological cells can be removed by oxidation with NaOCl solution [Georgieva, 2002]. After template decomposition the small decomposition products are released outside the polyelectrolyte shell without damage it. A very important feature of the obtained capsules is the possibility of opening and closing of the pores in their shell as a result of the variation of the solvent pH or composition. This property allows to fulfill the internal part of the capsule with some specific substances [Sukhorukov, 2005]. Different organic and inorganic molecules were inserted into the capsule volume, including enzymes for biocatalysis. Lvov at al. [Lvov, 2001b] reported the encapsulation of urease into 5 μm hollow capsules. Hollow polyelectrolyte capsules were fabricated by alternate adsorption of PSS and PAH onto melamine formaldehyde particles. The melamine formaldehyde particles were dissolved, and the obtained microcapsules were loaded with urease. The urease loading was done by varying the solvent composition. Confocal fluorescence microscopy reveled the close conformation of the capsules in a water solution containing labeled urease, after addition of ethanol (1:1 water ethanol mixture) penetration of urease into the capsules, indicating an open state of the capsule shell, was observed. When the loaded capsules were transferred into water, the polyion shells became closed for urease molecules which as a result were entrapped into the capsules volume. Urease encapsulated inside the polyion shell was found to preserve 13% of its activity as compared with free enzyme. This is a reasonable decrease due to the substrate diffusion into the capsules. After 5 days storage at 7 °C, encapsulated urease completely preserved its activity while free urease kept at the same conditions in aqueous solution lost 45% activity indicating that the polyion shell protected the encapsulated enzyme molecules. Enzyme capacity of hollow microcapsules (diameter ≅ 5μ)was found to be (105-107 molecules per particles) [Tiourina, 2001]. The microcapsule capacity was increased (108-109 molecules per particles) by the use of polyelectrolyte microspheres with filled interior [Balabushevich, 2003]. Microspheres of this kind were obtained by stepwise adsorption of
Laura Pastorino and Svetlana Erokhina
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natural polyelectolytes on melamine formaldehyde nuclei and subsequent partial decomposition of the nuclei under mild conditions. The partial decomposition of the nuclei resulted in a gel-like structure composed by the melamine formaldehyde residues complexed with the polyelectrolytes. Electrostatic and hydrophobic interactions were found to be responsible for the enzyme binding to the interior. The encapsulated peroxidase showed a high activity (57%), which remained stable for 12 months. Different protocols can be used for the enzyme incorporation. Zhu et al reported the encapsulation of glucose oxidase in diazoresin-based hollow polyelectrolyte capsules [Zhu, 2005]. The capsules were prepared via the alternate deposition of PSS and diazoresin on MnCO3 templates. After core dissolution, the capsules were exposed to a glucose oxidase solution for enzyme loading as such capsules were found to be permeable to large macromolecules. While still in the enzyme solution, the capsules were UV irradiated to crosslink the multilayer wall and reach the effective encapsulation. Encapsulated glucose oxidase revealed to retain 52.8% of the catalytic activity of the same amount of enzyme in solution. Also the shell properties can be tuned. The fabrication of mesoporous magnetic capsules was reported [Fang, 2002]. The capsules were fabricated by depositing magnetite nanoparticles and mesoporous silica nanoparticles onto polystyrene cores [Sadasivan, 2006] followed by core removing. Such high surface area composites display a high potentiality for biocatalytic applications where their magnetic properties could be used to separate them easily form the reaction products and for their self stirring. The wide range of nanobioreactors which can be fabricated using the different approaches described above makes this field of research extremely promising for the development of innovative biocatalytic systems.
Structure Characterization of LbL Based Nanobioreactors Prior to polyion multilayer formation on colloids, the coating procedure is elaborated onto a planar support and then transferred for the deposition on 3D templates. The deposition process on colloids can be then characterized qualitatively by electrophoretic mobility measurements. Namely, after each polyion coating deposition, the ζ-potential of the shell is calculated from the electrophoretic mobility (μ) using the Smoluchowski relationship [Smoluchowski, 1903]: ζ=μη/ε where η = viscosity of the solution ε = permittivity of the solution Successful deposition of a layer is determined by the observation of reversal of the charge on the colloid. The multilayer growth can be also evaluated by single particle light scattering, which allows the detection of the scattered light from a single particle at a given
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time [Lichtenfeld, 1995]. Confocal laser scanning microscopy (CLSM) can be used to image fluorescently labelled multilayer shell and to investigate the permeability of the capsule walls by monitoring the diffusion of fluorescent species into the capsule [Radtchenko, 2000; Lvov, 2001b]. AFM and transmission electron microscopy (TEM) are mainly used for the morphological characterization. AFM can be also used to collect valuable information on the mechanical properties of unloaded and loaded polyelectrolyte nanocapsules [Butt, 2005]. Recently the combination of CSLM, which gives information on the uniformity of film coverage, and flow cytometry, which rapidly measures the intensity of thousands of individual particles, have been proposed as a mean to characterization of polyelectrolyte multilayer films on particles [Johnston, 2006]. By comparing CLSM and flow cytometry data with QCM data on the same films assembled on planar supports it is possible to quantify the mass of polyelectrolytes deposited on the particles.
Multienzyme Films As already outlined, one of the most interesting features of the LbL technique is represented by its versatility which makes it possible to include a wide range of charged species within a multilayered structure with a predetermined architecture. This possibility can be used for the assembly of complex multienzyme films to be used in multistep biocatalytic processes for the production/degradation of complex species. The development of multistep biocatalytic routes in a single system is an important target for industrial biocatalysis [Schoemaker, 2003; Disawal, 2003] In 1996, Onda, Lvov at el deposited with LbL glucoamylase and glucose oxidase and 500 nm pore cellulose membrane (similar to filter paper) and got efficient two step vectorial bio/catalysis for transformation of starch to glucose, and then to sugar. It was demonstrated that only the proper two-enzyme architecture provides most efficient reaction outcome: first, glucoamylase layer for starch transformation, and then layer of glucose oxidase for glucose catalysis. The reaction was taken place in two-compartment reactor separated with enzyme modified membrane with 0.5 atm pressure applied to the first compartment. The highest catalytic production was reached when these two enzyme monolayers were separated by 10 nm interlayer of permeable polyelectrolytes [Onda, 1996a]. A final product separation was reached in such enzymatic reactor. This multi-enzyme catalysis in LbL multilayer was also developed for two-step glucose oxidase / peroxidase multilayer [Onda, 1996b]. The fabrication of multiprotein thin films on planar supports have been reported. A bienzyme system containing urease and arginase was fabricated and characterized. The multilayer structure was optimized for the improved catalytic activity by varying the number of enzyme layers and their position in the assembly. Also in this case the multilayers were shown to participate catalytically in a two step process for the decomposition of L-arginine to urea and subsequently to ammonia [Disawal, 2003]. Recently a novel strategy for multiprotein LbL self-assembly was reported. Namely, the LbL combination of the redox protein cytochrome c with the enzyme sulfite oxidase without use of any additional polymer was demonstrated [Dronov, 2008]. Electrostatic interactions between these two proteins, with
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rather separated isoelectric points during the assembly process from a low ionic strength buffer were found to be sufficient for the LbL layer deposition of both components. Mediator-free electron transfer of the enzyme within the film was achieved by coimmobilization of the enzyme and the redox protein from a mixture, rather than pure solutions. The electrocatalytic activity for sulfite oxidation generating catalytic current with a linear increase with the number of layers was demonstrated. Sequential enzymatic catalysis was demonstrated also for nanobiorectors. Multicomponent films containing glucose oxidase and horseradish peroxidase were prepared onto polystyrene particles and successive catalysis was shown [Caruso, 2000b]. The same enzymes were used also to demonstrate the potential application of polyelectrolyte multilayer microspheres as carriers of two functionally associated enzymes [Balabushevich, 2005]. Polyelectrolyte microspheres with filled interior [Balabushevich, 2003] were used to this aim. The LbL assembly of multiprotein systems for cascade bioreactions is a promising patway for the fabrication of a novel class of biocatalysts.
Conclusion In this chapter, two different approaches have been presented for the design and fabrication of complex multilayered structures containing enzyme monolayers, namely the LB and the LbL techniques. These techniques were found to be interesting for their application in the biocatalysis field as they make possible to control the immobilization process and consequently the biocatalyst properties at the nanoscale level. Moreover, the molecular dimension of the obtained biocatalysts makes economically feasible to work with expensive enzymes. As reported in this chapter, these techniques have been used for the immobilization of a variety of enzymes in complex architectures having a predetermined structure and function. It has been demonstrated that using these techniques it is possible to obtain highly efficient biocatalytic systems both in terms of residual catalytic activity and of functional stability. Interesting, the temporal stability of enzymes in such protected layers has been shown to be significantly higher even with respect to the free enzymes. Another advantage respect to traditional immobilization techniques is that the deposition of such structures can be carried out onto different supports, and in the case of the LbL technique it is possible to design and construct nano-bioreactors in which the enzyme molecules are deposited onto 3-D supports (e.g. nanoparticles) or are entrapped in the volume of a hollow nanocapsule. The versatility of these thin films techniques, makes possible to include different molecules and nano-objects in the desired position of the developed biocatalytic structures in order to impart specific properties. For example, conducting polymers have been used to facilitate electron transfer between the enzyme, substrate and product, and magnetic nanoparticles have been included in the layered shell of LbL based nano-bioreactors to impart them magnetic properties. The future prospects for such biocatalysts application and research appear promising, due to the steady developments in processing and characterization techniques. In conclusion it can be stated that the LB and LbL techniques can efficiently overcome some of the obstacles
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to a wider use of biocatalytic systems even at the industrial scale by developing next generation of biocatalysts.
Acknowledgments The authors would like to thank Dr V. Erokhin for valuable discussions and useful suggestions, and Konstantin Erokhin for his help in the figure preparation.
References Ai, H.; Gao, J. J Mater Sci. 2004, vol 39, 1429-1432. Agarwal, V. K.; Srivastava, V. K. J Appl Phys. 1973, vol 44, 2900-2901. Ai, H.; Lvov, Y.; Jones, S. Cell Biochemistry and Biophysics. 2003, v.39, 23-43. Aktsipetrov, O.A.; Mishina, E.D.; Murzina, T.V.; Novak, V.R.; Akhmediv, N.N. Thin Solid Films. 1995, vol 256, 176-181. Alcade, M.; Ferrer, M.; Plou, F. J.; Ballesteros, A. Trends Biotechnol. 2006, vol 24, 281-287. Alekseev M.E.; Myagkov, I.V. Colloid J. 1987, vol 5, 981-983. Aleu, J.; Bustillo, A. J.; Hernandez-Galan, R.; Collado, I. G. Curr Org Chem. 2006, vol 10, 2037-2054. Amador, S.M.; Myers, S.D.; Vodyanoy, V.; Neely, W.C. Biophys J. 1998, vol 75, 1783-1792. Angelova, A.; Vollhardt, D.; Ionov, R. J Phys Chem. 1996, vol 100, 10710-10720. Anicet, N.; Anne, A.; Moiroux, J.; Saveant, J.-M. J. Am. Chem. Soc. 1998, vol. 120, 71157116. Antipov, A. A.; Shchukin, D.; Fedutik, Y.; Petrov, A. I.; Sukhorukov, G. B.; Mohwald, H. Colloids Surf, A. 2003, vol 224, 175-183. Anzai, J.; Lee, S.; Osa, T. Macromol Chem Rapid Commun. 1989, vol 10, 167-170. Ariga, K.; Lvov, Y.; Kunitake, T. J Am Chem Soc. 1997, vol 119, 2224-2231. Babakov, A. V.; Terekhov, O. P.; Yanenko, A. S. Biochim Biophys Acta. 1979, vol 547, 438446. Bai, Y.; Zhao, S.; Zhang, K.; Sun, C. Colloids Surf, A. 2006, vol 281, 105-112. Balabushevich, N. G. ; Tiourina, O. P.; Volodkin, D.; Larionova, N.; Sukhorukov, G. B. Biomacromolecules. 2003, vol 4, 1191-1197. Balabushevich, N. G.; Sukhorukov, G. B.; Larionova, N. I. Macromol Rapid Commun. 2005, vol 26, 1168-1172. Barmin, A. V.; Eremenko, A.; Kurochkin, I. N.; Moskvitina, T. A. Biotechnol Bioengineer. 1994, vol 245, 206-210. Berzina, T. S.; Piras, L.; Troitsky, V. I. Thin Solid Films. 1998, vol 327-329, 621-626. Bianco, I. D.; Fidelio, G. D.; Maggio, B. Biochem J. 1989, vol 258, 95-99. Blinov, L. M.; Dubinin, N. V.; Mikhnev, L. V.; Yudin, S. G. Thin Solid Films. 1984, vol 120, 161-170. Blodgett, K. B. J Am Chem Soc. 1934, vol 56, 495. Blodgett, K. B. J Am Chem Soc. 1935, vol 57, 1007-1022.
38
Laura Pastorino and Svetlana Erokhina
Blodgett, K. B.; Langmuir, I. Phys Rev. 1937, vol 51, 964-982. Bommarus, A. S.; Polizzi, K. M. Chem Eng Sci. 2006, vol 61, 1004–1016. Brynda, E.; Pacherník, J.; Houska, M.; Pientka, Z.; Dvořák, P. Langmuir. 2005, vol 21, 78777883. Butt, H. J.; Cappella, B.; Kappl, M. Surf Sci Rep. 2005, vol. 59, 1–152. Cao, L. Curr Opin Chem Biol. 2005, vol 9, 217-226. Caruso, F.; Caruso, R.; Mohwald, H. Science. 1998, vol 282, 1111-1114. Caruso, F.; Fiedler, H.; Haage, K. Colloids Surf, A. 2000a, vol 169, 287-293. Caruso, F.; Möhwald, H. J Am Chem Soc. 1999, vol 121, 6039-6046. Caruso, F.; Niikura, K.; Furlong, D.; Okahata, Y. Langmuir. 1997, vol 13, 3427-3433. Caruso, F.; Schüler, C. Langmuir. 2000b, vol 16, 9595-9603. Caruso, F.; Trau, D.; Mohwald, H.; Renneberg, R. Langmuir. 2000c, vol 16, 1485-1488. Caseli, L.; Moraes, M. L.; Zucolotto, V.; Ferreira, M.; Nobre, T. M.; Zaniquelli, M. E. D.; Filho, U. P. R.; Oliveira Jr., O. N. Langmuir. 2006, vol 22, 8501-8508. Cassier, T.; Lowack, K.; Decher, G. Supramol Sci. 1998, vol 5, 309-315. Castro, L. B. R.; Silva, F. F.; Carmona-Ribeiro, A. M.; Kappl, M.; Petri, D. F. S. J Phys Chem B. 2007, vol 111, 8520-8526. Chasovnikova, L. V.; Formaziuk, V. E.; Sergienko, V. I.; Kokriakov, V. N. Biokhimiia. 1992, vol 57, 97-102. Chi, L. F.; Johnston, R. R.; Ringsdorf, H. J Am Chem Soc. 1987, vol 109, 788-796. Chibata, I.; Tosa, T.; Sato, T. J Mol Catal. 1986, vol 37, 1-24. Choi, J. W.; Kim, Y. K.; Lee, I. H.; Min, J.; Lee, W. H. Biosens Bioelectron. 2001, vol 16, 937-943. Choi, J. W.; Min, J. H.; Jung, J. W.; Rhee, H. W.; Lee, W. H. Mol Cryst Liq Cryst. 1997, vol 294, 451-454. Colacicco, G. Nature. 1971, vol 233, 202-204. Coward-Kelly, G.; Ruizhen Chen, R. Biotechnol Prog. 2007, vol 23, 52-54. Cristofolini, L.; Berzina, T.; Erokhina, S.; Konovalov, O.; Erokhin, V. Biomacromolecules. 2007, vol 8, 2270-2275. Cullison, J. K.; Hawkridge, F. M.; Nakashima, N.; Yoshikawa, S. Langmuir. 1994, vol 10, 877-883. Dai, Z.; Wilson, J. T.; Chaikof, E. L. Mater Sci Eng, C. 2007, vol 27, 402-408. Danilov, V. S.; Sitkovskii, M. V.; Kozlov, Y. P.; Chukhrai, E. S.; Kamyshnyi, A. S. Biofizika. 1975, vol 20, 441-444. Davies, D. E.; Critchlow, M.; Grainger, D. W.; Reichert, A.; Ringsdorf, H.; Lloyd, J. B. Biochim Biophys Acta. 1991, vol 1084, 29-34. Decher, G. Science. 1997, vol 227, 1232-1237. Decher, G.; Hong, J. D. Makromol Chem Macromol Symp. 1991, vol 46, 321-327. den Engelsen, D.; de Koning, B. R.; J Chem Soc Faraday Trans. 1974, vol 70, 2100-2112. Di Natale, C.; Goletti, C.; Paolesse, R.; Drago, M.; Macagnano, A.; Mantini, A. Troitsky, V. I.; Berzina, T. S.; Cocco, M.; D’Amico, A. Sens Actuators, B. 1999, vol 57, 183-187. Disawal, S.; Qiu, J.; Elmore, B. B.; Lvov, Y. M. Colloids Surf, B. 2003, vol 32, 145-156. Donath, E.; Sukhorukov, G. B.; Caruso, F.; Davis, S. A.; Mohwald, H. Angew Chem Int Ed. 1998, vol 37, 2202-2205
Nanobiocatalytic Systems: Thin Films of Enzymes
39
Dronov, R.; Kurth, D.; Möhwald, H.; Spricigo, R.; Leimkühler, S.; Wollenberge, U.; Rajagopalan, K.; Scheller, F.; Lisdat, F. J Am Chem Soc. 2008, vol 130, 1122 -1123. Ducharme, D.; Salesse, C.; Leblanc, R. M. Thin Solid Films. 1985, vol 132, 83-90. Dudnik, V.; Sukhorukov, G. B.; Radtchenko, I. L.; Mohwald, H. Macromolecules, 2001, vol 34, 2329–2334. Dziri, L.; Desbat, B.; Leblanc, R. M. J Am Chem Soc. 1999, vol 121, 9618-9625. Edmiston, P. L.; Saavedra, S. S. Biophys. J. 1998, vol 74, 1006. Eremenko, A. V.; Mineeva, M. F.; Chernov, S. F.; Kurochkin, I. N. Bull. Experimentalnoiy Biologii. 1991, vol 9, 246-248. Erokhin, V. Langmuir-Blodgett Films of Biological Molecules, in Handbook of Thin Fims Materials (N.S. Nalwa, ed.), Academic Press, San Diego, 2002, vol1, pp. 523-558. Erokhin, V. Langmuir-Blodgett multilayers of proteins, in Protein Architecture: Interfacing Molecular Assemblies and Immobilization Biotechnology (Y. Lvov and H. Möhwald, eds.), Marcel Dekker, Inc., New York. 2000, pp. 99-124. Erokhin, V.; Lvov, Y.; Mogilevsky, L.; Zozulin, A.; Ilyin, E. Thin Solid Films. 1989, vol 178, 153-158. Erokhina, S.; Erokhin, V., Nicolini, C. Colloids and Surfaces A. 2002, vol. 198-200, 645-650. Erokhina, S.; Berzina, T.; Cristofolini, L.; Konovalov, O.; Erokhin, V.; Fontana, M. P. Langmuir. 2007, vol 23, 4414-4420. Estrela-Lopis, I.; Brezesinski, G.; Möhwald, H. Biophys J. 2001, vol 80, 749-754. Facci, P.; Erokhin, V.; Nicolini, C.Thin Solid Films. 1993, vol 230, 86-89. Facci, P.; Erokhin, V.; Tronin, A.; Nicolini, C. J. Phys. Chem. 1994, vol. 98, 13323-13327. Faergemand, M.; Murray, B. S.; Dickinson, E. J Agric Food Chem. 1997, vol 45, 2514-2519. Fang, M.; Grant, P.; McShane, M.; Sukhorukov, G.; Golub, V.; Lvov, Y. Langmuir. 2002, vol 18, 6338-6344. Feachem, C. G. P.; Tronstad, L. Proc Roy Soc London Ser A. 1934, vol 145, 127-135. Fromherz, P. Nature. 1971, vol 231, 267-268. Fromherz, P.; Marcheva, D. FEBS Lett. 1975, vol 49, 329-333. Frutos, A. G.; Corn, R. M. Anal Chem. 1998, vol 70, 449–455. Gaines, Jr. G.L. Insoluble Monolayers at Liquid-Gas Interface, Wiley-Interscience, New York, 1956. Geogieva, R.; Moya, S.; Hin, M.; Mitlohner, R.; Donath, E.; Kiesewetter, H.; Mohwald, H.; Baumler, H. Biomacromolecules. 2002, vol 3, 517-524. George, S. P.; Gole, A. M.; Sastry, M.; Rao, M. B. Langmuir. 2002, vol 18, 9494-9451. Gidalevitz, D.; Huang, Z.; Rice, S.vA. Biophys J. 1999, vol 76, 2797-2802. Giocondi, M. C.; Besson, F.; Dosset, P.; Milhiet, P. E.; Le Grimellec, C. Langmuir. 2007, vol 23, 9358-9364. Girard-Egrot, A. P.; Morelis, R. M.; Coulet, P. R. Langmuir. 1997, vol 13, 6540-6546. Girard-Egrot, A. P.; Morelis, R. M.; Coulet, P. R. Langmuir. 1998, vol 14, 476-482. Girard-Egrot, A. P.; Morélis, R. M.; Coulet, P. R. Thin Solid Films. 1997, vol 232, 282-289. Godoy, S.; Leca-Bouvier, B.; Boullanger, P.; Blum, L. J.; Girard-Egrot, A.P. Sens Actuators B. 2005, vol 107, 82-87. Gole, A.; Dash, C.; Mandale, A. B.; Rao, M.; Sastry, M. Anal Chem. 2000, vol 72, 43014309.
40
Laura Pastorino and Svetlana Erokhina
Grandbois, M.; Desbat, B.; Blaudez, D.; Salesse, C. Langmuir. 1999, vol 15, 6594-6597. Hamlin, R. E.; Dayton, T. L.; Johnson, L. E.; Johal, M. S. Langmuir. 2007, vol 23, 4432 4437. Harris, J. R.; Holzenburg, A. J Struct Biol. 1995, vol 115, 102-112. Hartel, S.; Fanani, M. L.; Maggio, B. Biophys J. 2005, vol 88, 287-304. Hartmeier, W. Immobilized Biocatalysts, Springer-Verlag: Berlin Heidelberg, DE, 1988. Helm, C. A.; Möhwald, H.; Kjaer, K.; Als-Nielsen, J Europhys Lett. 1987, vol 4, 697-703. Hernández-Martín, E.; Otero, C. Bioresour Technol. 2008, vol 99, 277-286. Hou, Y.; Jaffrezic-Renault, N.; Zhang, A.; Wan, J.; Errachid, A.; Chovelon, J. M. Sens Actuators B. 2002, vol 86, 143-149. Hourdou, M. L.; Besson, F. Biotechnol Techniques. 1995, vol 9, 643-648. Huie, J. C. Smart Mater Struct. 2003, vol 12, 264-271. Iler, R. J Colloid Interface Sci. 1966, vol 21, 569-594. Inbar, L.; Miller, I. R. Biochim Biophys Acta. 1976, vol 452, 421-430. Ivanova, T. Z.; Raneva, V.; Panaiotov, I.; Verger, R. Colloid Polym. Sci. 1993, vol 271, 290297. Ivanova, T.; Malzert, A.; Boury, F.; Proust, J. E.; Verger, I. Colloids Surf B. 2003, vol 32, 307-320. Jensen, M. M. O.; Jensen, T. R.; Kjaer, K.; Bjornholm, T.; Mouritsen, O. G.; Peters, G. H. Biophys J. 2002, vol 83, 98-111. Jin, W., Xiangyang, S.; Caruso, F. J Am Chem Soc. 2001, vol 123, 8121-8122. Johnston, A. P. R.; Zelikin, A. N.; Lee, L.; Caruso, F. Anal Chem. 2006, vol 78, 5913-5919. Kaganer, V. M.; Brezesinski, G.; Möhwald, H.; Howes, P. B.; Kjaer, K. Phys Rev E. 1999, vol 59, 2141-2152. Karnati, C.; Du, H.; Ji, F.; Lvov, Y.; Chen, W. Biosens Bioelectron. 2007, vol 22, 2636-2642. Katchalsky-Katzir, E. Trends Biotechnol. 1993, vol 11, 471-478. Kato, T.; J J. Appl Phys. 1987, vol 26, L1377-L1380. Kim, J.; Grate, J. W.; Wang, P. Chem Eng Sci. 2006b, vol 61, 1017-1026. Kim, J.; Jia, H.; Wang, P. Biotechnol Adv. 2006a, vol 24, 296– 308. Kim, M. W.; Sauer, B. B.; Yu, H.; Yazdanian, M.; Zografi, G. Langmuir. 1990, vol 6, 236240. Kjaer, K.; Als-Nielsen, J.; Helm, C. A.; Tippmann-Krayer, P.; Möhwald, H. Thin Solid Films. 1988, vol 159, 17-28. Knoll, W. Curr Opin Colloid Interf Sci. 1996, vol 1, 137–143. Koeller, K. M.; Wong, C. H. Nature. 2001, vol 409, 232-240. Kong, J.; Lu, Z.; Lvov, Y.; Desamero, R.; Frank, H.; Rusling, J. J Am Chem Soc., 1998, vol 120, 7371-7372. Kristensen, D.; Nylander, T.; Rasmussen, J. T.; Paulsson, M.; Birdi, K. S. Colloids Surf A. 1998, vol 143, 221-231. Kuhn, H. Chem Phys Lipids. 1972, vol 8, 401-404. Kuhn, H. J Chem Phys. 1970, vol 53, 101-108. Kuhn, H. Pure Appl Chem. 1979, vol 51, 341-352. Lagocki, J. W.; Boyd, N. M.; Law, J. H.; Kézdy, F. J. J Am Chem Soc. 1970, vol 92, 29232924.
Nanobiocatalytic Systems: Thin Films of Enzymes
41
Langmuir, I. J Am Chem Soc. 1917, vol 39, 1848-1906. Langmuir, I.; Schaefer, V. J. J Am Chem Soc. 1938, vol 60, 1351-1360. Lebeau, L.; Olland, S.; Oudet, P.; Mioskowski, C. Chem Phys Lipids. 1992, vol 62, 93-105. Lebeau, L.; Regnier, E.; Schultz, P.; Wang, J. C.; Mioskowski, C.; Oudet, P. FEBS Lett. 1990, vol 267, 38-42. Lee, S.; Anzai, J.; Osa, T. Sens Actuators B. 1993, vol 12, 153-158. Lee, Y. L.; Lin, J. Y.; Lee, S. Langmuir. 2007, vol 23, 2042-2051. Li, Q.; Quinn, J. F.; Wang, Y.; Caruso, F. Chem Mater. 2006, vol 18, 5480-5485. Liang, Z.; Wang, C.; Tong, Z.; Ye, W.; Ye, S. React Funct Polym. 2005, vol 63, 85-94. Lichtenfeld, H.; Knapschinsky, L.; Sonntag, H.; Shilov, V. Colloids Surf, A. 1995, vol 104, 313-320. Lösche, M.; Möhwald, H. Rev Sci Instrum. 1984, vol 55, 1968-1972. Lu, Z.; Li, C. M.; Zhou, Q.; Bao, Q. L.; Cui, X. J Colloid Interface Sci. 2007, vol 314, 80-88. Lvov, Y. In Encyclopedia of Surface and Colloid Science; Hubbard, A.; Marcel Dekker: New York, NY, 2002, pp 321-349. Lvov, Y. In Protein Architecture: Interfacial Molecular Assembly and Immobilization Biotechnology; Lvov, Y.; Möhwald, H.; Marcel Dekker: New York, NY, 2000, pp 125136. Lvov, Y.; Lu, Z.; Zu, X.; Schenkman, J.; Rusling, J. J Am Chem Soc. 1998, vol 120, 40734080. Lvov, Y.; Ariga, K.; Ichinose, I.; Kunitake, T. J Am Chem Soc. 1995, vol 117, 6117-6123. Lvov, Y.; Caruso, F. Analyt Chem. 2001a, vol 73, 4212-4217. Lvov, Y.; Decher, G. Crystallogr Rep. 1994, vol 39, 628-647. Lvov, Y.; Feigin, L. Studia Biophysica. 1986, vol 112, 221-224. Lvov, Y; Antipov, A. A.; Mamedov, A.; Mohwald, H.; Sukhorukov, G. B. Nanoletters. 2001b, vol 1, 125-128. Majewski, J.; Kuhl, T. L.; Kjaer, K.; Gerstenberg, M. C.; Als-Nielsen, J.; Israelachvili, J. N.; Smith, G. S. J Am Chem Soc. 1998, vol 120, 1469-1473. Maksymiw, R.; Nitsch, W. J Colloid Interface Sci. 1991, vol 147, 67-77. Marron-Brignone, L.; Morélis, R. M.; Chauvet, J. P.; Coulet, P. R. Langmuir. 2000, vol 16, 498-503. Marron-Brignone, L.; Morelis, R. M.; Coulet, P. R. Langmuir. 1996, vol 12, 5674-5680. Mateo, C.; Palomo, J. M.; Fernandez-Lorente, G.; Guisan, J. M.; Fernandez-Lafuente, R. Enzyme Microb Technol. 2007, vol 40, 1451-1463. Matsuno, H.; Niikura, K.; Okahata, Y. Chem. Eur. J. 2001, vol 7, 3305-3312. Matsuo, Y.; Ijiro, K.; Shimomura, M. Colloids and Surfaces B. 2005, vol 40, 123-126. Mello, S. V.; Coutures, C.; Leblanc, R. M.; Cheng, T. C.; Rastogi, V. K.; DeFrank, J. J. Talanta. 2001, vol 55, 881-887. Mello, S. V.; Mabrouki, M.; Cao, X.; Leblanc, R. L.; Cheng, T. C.; DeFrank, J. J. Biomacromolecules. 2003, vol 4, 968-973. Mirsky, V. M. Chem Phys Lipids, 1994, vol 70, 75-81. Mizushina, Y.; Sagisaka, M.; Sakai, H.; Abe, M.; Sakaguchi, K. Biochim Biophys Acta. 2000, vol 1486, 211-218. Mohwald, H. Colloids Surf, A. 2000, vol 17, 25–31.
42
Laura Pastorino and Svetlana Erokhina
Morandat, S.; Bortolato, M.; Nicol, F.; Arthur, J. R.; Chauvet, J. P.; Roux, B. Colloids Surf B. 2004, vol 35, 99-105. Morgan, H.; Taylor, D .M. Biosens Bioelectron. 1992, vol 7, 405-410. Nagahori, N.; Niikura, K.; Sadamoto, R.; Taniguchi, M.; Yamagishi, A.; Monde, K.; Nishimura, S. I. Adv Synthesis and Catalysis. 2003, vol 345, 729-734. Nakamura, C.; Mizutani, W.; Lantz, M. A.; Noda, K.; Zorin, N.A.; Miyake, J. Supramol Sci. 1998, vol 5, 639-642. Nielsen, L. K.; Balashev, K.; Callisen, T. H.; Bjornholm, T. Biophys J. 2002, vol 83, 26172624. Nitsch, W.; Maksymiw, R. Colloid Polymer Sci. 1990, vol 268, 452-459. Noda, K.; Zorin, N. A.; Nakamura, C.; Miyake, M.; Gogotov, I. N.; Asada, Y.; Akutsu, H.; Miyake, J. Thin Solid Films. 1998, vol 327, 639-642. Okahata, Y.; Tsuruta, T.; Ijiro, K.; Ariga, K. Thin Solid Films. 1989, vol 180, 65-72. Okawa, Y.; Tsuzuki, H.; Yoshida, S.; Watanabe, T. Anal. Sci. 1989, vol. 5, 507. Oliveira, Jr., O. N.; Taylor, D. M.; Stirling, C. J. M.; Tripathi, S.; Guo, B. Z. Langmuir. 1992, vol 8, 1619-1626. Onda, M.; Lvov, Y.; Ariga, K.; Kunitake, T. J Ferment Bioeng. 1996a, vol 82, 502-506. Onda, M.; Lvov, Y.; Ariga, K., Biotechnol Bioeng., 1996b, vol. 51, 163-166. Overbeck, G. A.; Hönig, D.; Wolhaus, L.; Gnade, M.; Möbius, D. Thin Solid Films. 1994, vol 242, 26-32. Overbeck, G. A.; Möbius, D. J Phys Chem. 1993, vol 97, 7999-8004. Pachence, J. M.; Blasie, J. K. Biochem J. 1991, vol 59, 894-900. Paddeu, S.; Erokhin, V.; Nicolini, C. Thin Solid Films. 1996, vol 284, 854-858. Pal, P.; Nandi, D.; Misra, T. N. Thin Solid Films. 1994, vol 239, 138-143. Palomba, S.; Berovic, N.; Palmer, R. E. Langmuir. 2006, vol 22, 5451-5454. Pastorino, L.; Berzina, T. S.; Troitsky, V. I.; Bernasconi, E.; Nicolini, C. Colloids Surf B. 2002, vol 23, 289-293. Pastorino, L.; Disawal, S.; Nicolini, C.; Lvov, Y. M.; Erokhin, V. V. Biotechnol Bioeng. 2003, 84, 286-291. Pastorino, L.; Nicolini, C. Mater Sci Engineer C. 2002, vol 22, 419-422. Patel, D. S.; Aithal, R. K.; Krishna, G.; Lvov, Y. M.; Tien, M.; Kuila, D. Colloids Surf, B. 2005, vol 43, 13-19. Peschke, J.; Möhwald, H. Colloids Surf. 1987, vol 27, 305-323. Peters, J.; Fromherz, P. Biochim Biophys Acta. 1975, vol 394, 111-119. Petersen, M. Curr Opin Biotechnol. 1999, vol 10, 593-594. Peterson, I. R. Phys Scripta. 1992, T45, 245-247. Petrigliano, A.; Tronin, A.; Nicolini, C. Thin Solid Films. 1996, vol 284, 752-756. Petrov, A. I.; Volodkin, D. V.; Sukhorukov, G. B. Biotechnol Prog. 2005, vol 21, 918-925. Pollard, D. J.; Woodley, J M Trends Biotechnol. 2006, vol 25, 66-73. Prokop, L. A.; Strongin, R. M.; Smith III, A. B.; Blasie, J. K.; Peticolas, L. J.; Bean, J. C. Biophys J. 1995, vol 70, 2131-2143. Qian, D. J.; Nakamura, C.; Zorin, N.; Miyake, J. Colloids Surf A. 2002, vol 198, 663– 669. Radtchenko, I. L.; Sukhorukov, G. B.; Leporatti, S.; Khomutov, G. B.; Donath E.; Möhwald, H. J Colloid Interface Sci. 2000, vol 230, 272-280.
Nanobiocatalytic Systems: Thin Films of Enzymes
43
Radtchenko, I. L.; Sukhorukov, G. B.; Mohwald, H. Colloids Surf, A. 2002, vol 202, 127– 133. Rajdev, P.; Chatteji, D. Langmuir. 2007, vol 23, 2037-2041. Rao, C. S.; Damodaran, S. Colloids Surf A. 2005, vol 45, 49-55. Rashid, U.; Anwar, F. Fuel. 2008, vol 87, 265-273. Razumas, V.; Nylander, T.; MacRitchie, F. J Colloid Interface Sci. 1996, vol 178, 303-308. Rinuy, J.; Brevet, P. F.; Girault, H. H. Biophys J. 1999, vol 77, 3350-3355. Roberts, G. G. Ferroelectrics. 1989, vol 91, 21-38. Roberts, G.G. Langmuir-Blodgett Films. Plenum Press, New York, 1990. Rodríguez Couto, S.; Toca Herrera, J. L. Biotechnol Adv. 2006, vol 24, 500–513. Rosilio, V.; Boissonnade, M.-M.; Zhang, J.; Jiang, L.; Baszkin, A. Langmuir. 1997, vol. 13, 4669-4675. Rubin-Pitel, S. B.; Zhao, H. Comb Chem High Throughput Screening. 2006, vol 9, 247-257. Rusling, J. Am. Chem. Soc., 1998, v.120, 7371-7372. Sadasivan, S.; Sukhorukov, G. B. J Colloid Interface Sci. 2006, vol 304, 437-441. Salamon, Z.; Hazzard, J. T.; Tollin, G. Proc Natl Acad Sci. USA. 1993, vol 90, 6420-6423. Salditt, T.; Schuber, U. S. Rev Mol Biotechnol. 2002, vol 90, 55-70. Sano, M.; Lvov, Y.; Kunitake, T. Annu Rev Mater Sci. 1996, vol 26, 153-187. Sauerbrey, G. Z. Z Phys. 1959, vol 155, 206-218. Schäffer, C.; Novotny, R.; Küpcü, S.; Zayni, S.; Scheberl, A.; Friedmann, J.; Sleytr, U.B.; Messner, P. Small. 2007, vol 9, 1549-1559. Schmid, A.; Dordick, J. S.; Hauer, B.; Kiener, A.; Wubbolts, M.; Witholt, M. Nature. 2001, vol 409, 258-268. Schoemaker, H. E.; Mink, D.; Wubbolts, M. G. Science. 2003, vol 299, 1694-1697. Schonhoff, M. Curr Opin Colloid Interface Sci. 2003, vol 8, 86-95. Schuler, C.; Caruso, F. Macromol Rapid Commun. 2000, vol 21, 750–753. Schultz, P.; Vautier, D.; Richert, L.; Jessel, N.; Haikel, Y; Schaaf, P; Voegel, J. C.; Ogier, J.; Debry, J. Biomaterials. 2005, vol 26, 2621-2630. Schwartz, D. K.; Knobler, C .M. J Phys Chem. 1993, vol 97, 8849-8851. Seeman, N. C. Int J Nanotech. 2005, vol 2, 348-370. Shchukin, D. G.; Patel, A. A.; Sukhorukov, G. B.; Lvov, Y. M. J Am Chem Soc. 2004, vol 126, 3374-3375. Sheldon, R.A.; van Rantwijk, F. Aust J Chem. 2004, vol 57, 281-289. Shen, B. W.; Tsao, F. H.; Law, H. J.; Kézdy, F.J. J Am Chem Soc. 1975, vol 97, 1205-1208. Shimomura, M.; Sawadaishi, T. Curr Opin Colloid Interface Sci. 2001, vol 6, 11 16. Shutava, T.; Kommireddy, D.; Lvov, Y. J Am Chem Soc. 2006, vol 128, 9926-9934. Shutava, T.; Zheng, Z.; John, V.; Lvov, Y. Biomacromolecules. 2004, vol 5, 914-921. Singhal, R.; Gambhir, A.; Pandey, M. K.; Annapoorni, S.; Malhotra, B. D. Biosens Bioelectron. 2002, vol 17, 697-703. Slotte, J. P.; Ostman, A. L. Biochim Biophys Acta. 1993, vol 1145, 243-249. Slotte, J.P. Biochim Biophys Acta. 1992, vol 1124, 23-28. Sobotka, H.; Bloch, E. J Phys Chem. 1941, vol 45, 9-19. Stein, E. W.; McShein, M. J. IEEE T Nonobiosci. 2003, vol 2, 133-137.
44
Laura Pastorino and Svetlana Erokhina
Sukhorukov, G. B. in Novel Methods to Study Interfacial layers; Mobius, D.; Miller, R.; Studies in Interface Science, 11;Elselvier, 2001; Vol. 11, pp 383-414. Sukhorukov, G.; Fery, A.; Möhwald, H. Progr Polym Sci. 2005, vol 30, 885-897. Sun, S.; Ho-Si, P. H.; Harrison, D. J. Langmuir. 1991, vol 7, 727-737. Surplice, N. A.; D’Arcy, R. J. J Phys E. 1970, vol 3, 477-482. Szabo´, T.; Szekeres, M.; Dékány,I.; Jackers,C.; De Feyter, S.; Johnston, C. T.; Schoonheydt, R. A. J Phys Chem C. 2007, vol 111, 12730-12740 Tang, J. L.; Jiang, J. G.; Song, Y. H.; Peng, Z. Q.; Wu, Z. Y.; Dong, S. J.; Wang, E. K. Chem Phys Lipids. 2002, vol 120, 119-129. Tiourina, O.; Antipov, A.; Sukhorukov, G. B.;Larinova, N; Lvov, Y.; Mohwald, H. Macromol Biosci. 2001, vol 1, 209-214. Tredgold R. H.; Elgamal M. Biochim Biophys Acta. 1979, vol 555, 381-387. Troitsky, V. I.; Sartore, M.; Berzina, T. S.; Nardelli, D.; Nicolini, C. Rev Sci Instrum. 1996, vol 67, 4216-4223. Tsao, M. W.; Fischr, T. M.; Knobler, C. M. Langmuir. 1995, vol 11, 3184-3188. Ulman, A. An Introduction to Ultrathin Organic Films: from Langmuir-Blodgett to Selfassembly. Academic press: Boston, MA, 1991; pp 1-440. Uzgiris, E. E. Biochem J. 1987, vol 242, 293-296. van Beilen, J. B.; Zhi, L. Curr Opin Biotechnol. 2002, vol 13, 338-344. Vinod, V. P.; Phadtare, S.; Joshi, H. M.; Sastry, M.; Rao, M. J Nanosci Nanotechnol. 2007, vol 7, 2767-2773. Voigt, U.; Khrenov, V.; Tauer, K.; Hahn, M.; Jaeger, W.; von Klitzing, R. J Phys Condens Matter. 2003, vol 15, 213–218. von Smoluckowski, M. Bulletin International de l’Académie des Sciences de Cracovie. 1903, vol 8, 182–199. Wang, C.; Li, C.; Ji, X.; Orbulescu, J.; Xu, J.; Leblanc, R. M. Langmuir. 2006, vol 22, 22002204. Wang, P. Curr Opin Biotechnol. 2006, vol 17, 574–579. Wang, X.; Zheng, S.; He, Q.; Brezesinski, G.; Möhwald, H.; Li, J. Langmuir. 2005, vol 20, 1051-1054. Watanabe, N.; Ohnuki, H.; Saiki, T.; Endo, H.; Izumi, M.; Imakubo, T. Synth Met. 2005, vol 153, 25-28. Whiteley, C. G.; Lee, D. J. Enzyme Microb Technol. 2006, vol 38, 291-316. Wilhelmy, L. Ann. Phys. Chem. 1863, vol 119, 177. Wu, B. J.; Else, P. L.; Storlien, L. H.; Hulbert, A. J. J Exp Biol. 2001, vol 204, 4271-4280. Wu, L. Q.; Payne, G. F. Trends Biotechnol. 2004, vol 22, 593-599. Wu, Y.; Teng, Y.; Li, Z.; Liao, X.; Luo, Y. Soil Biol Biochem. 2008, vol 40, 789-796. Xing, Q.; Eadula, S.; Lvov, Y. Biomacromolecules. 2007, vol 8, 1987-1991. Yasutake, M.; Aoki, D.; Fujihira, M. Thin Solid Films. 1996, vol 273, 279-283. Yu, A. M.; Liang, Z. J.; Caruso, F. Chem Mat. 2005, vol 17, 171-175. Zaitsev, S. Y. Colloids Surf B. 1993, vol 75, 211-216. Zayats, M.; Katz, E.; Willner, I. J Am Chem Soc. 2002, vol 124, 2120-2121. Zhang, J.; Rosilio, V.; Goldmann, M.; Boissonnade, M. M.; Baszkin, A. Langmuir. 2000, vol 16, 1226-1232.
Nanobiocatalytic Systems: Thin Films of Enzymes
45
Zhou, D.; Kang, D. J. Int J Nanotech. 2005, vol 2, 440-468. Zhu, H.; McShane, M. Langmuir. 2005, vol 21, 424-430.
Reviewed by Prof. Yuri Lvov, Institute for Micromanufacturing, Louisiana Tech University, Ruston Louisiana.
In: Biocatalysis Research Progress Editors: F. H. Romano, A. Russo
ISBN: 978-1-60456-619-2 © 2008 Nova Science Publishers, Inc.
Chapter II
Bioprocesses for the Synthesis of Nucleosides and Nucleotides Marco Terreni,* Daniela Ubiali, Teodora Bavaro, Davide A. Cecchini, Immacolata Serra and Massimo Pregnolato Italian Biocatalysis Center, PBL Dipartimento di Chimica Farmaceutica, via Taramelli 12, Università degli Studi, I-27100 Pavia, Italy Phone: +39-0382-987265, Fax: +39-0382-422975 E-mail:
[email protected]
Abstract Modified nucleosides and nucleotides are routinely used as citotoxic or antiviral drugs as well as immunosuppressive agents. The therapeutic activity of these compounds is due to their ability to act as antimetabolites in the RNA and DNA synthesis. Nucleosides have traditionally been prepared by various chemical methods, involving multi-step chemical procedures which are plagued by low yields and the formation of undesired by-products. These drawbacks strongly reduce the performances of the processes used for the synthesis of unnatural nucleosides and nucleotides, in terms of yields, purity of the final product, costs and environmental impact. Enzymatic syntheses have been shown to be an advantageous alternative to chemical methods due to the high selectivity of the enzymes, the mild reaction conditions and the overall simplicity of the approach. These bioprocesses can overcome the need of specific protecting groups often required on the heterocyclic base and/or on the sugar residue in the glycosylation reaction as well as in the modification of naturally occurring nucleosides. Consequently, these processes can be very competitive in terms of costs, allowing high quality products to be obtained in high yields.
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Marco Terreni, Daniela Ubiali, Teodora Bavaro, et al. Glycosyl-transferring enzymes (e.g. nucleoside phosphorylases, Ndeoxyribosyltransferases) have been used in the synthesis of nucleosides by mediating the enantioselective transfer of glycosyl residues to acceptor bases. Recent reports involving glycosyl-transferring enzymes are reviewed herein. In this context, the use of isolated deaminases to obtain, respectively, cytidine derivatives from uridine, or guanosine derivatives from adenosine is also considered. Moreover, hydrolases (e.g. lipases, esterases) will be described in carrying out regio-controlled manipulations on the carbohydrate moiety. Finally, the enzyme-mediated synthesis of nucleotides through regioselective phosphorylation of the parent nucleosides catalyzed by kinases and phosphatases are discussed. This review will also focus on the importance of the enzyme selection and the design of the final biocatalyst thereof, particularly by tailor-made immobilization of the protein on solid support, with the aim to have an active, stable and reusable biocatalyst for preparative purposes.
1. Introduction Modified nucleosides and nucleotides belong to an important class of therapeutic agents with antiviral and anti tumour activity and are key raw materials for the preparation oligonucleotides [1]. These compounds are routinely synthesized by multi-step chemical procedures [2] which are often plagued by the formation of undesired by-products and low overall yields [3]. When nucleoside and nucleotides are intended for human use, impurities content must be strictly controlled. To pursue this goal, chemical procedures do require several purification steps to guarantee the quality standard defined by regulatory authorities for the use of active pharmaceutical ingredients (API). Consequently, the overall performances of the chemical processes may be negatively affected in terms of yield, costs and environmental impact. The quality of a pharmaceutical product is very important and regulation is becoming stricter, particularly in terms of impurities control. The EMEA’s Committee for Medicinal Products for Human Use (CHMP) provides scientific guidelines for marketing-authorization applications related to medicinal products. Guidelines are intended to provide a basis for practical harmonization of the manner in which the EU member states interpret and apply the detailed requirements for the demonstration of quality, safety and efficacy contained in the Community directives. According to EMEA guidelines, strict limits for impurities in new drugs are assessed: any impurity should be below 0.10%, otherwise, complete characterization and toxicological evaluation of each impurity exceeding this limit is required [4]. For generic products (already on the market), fine chemical industries develop alternative synthetic routes. In this case, new unknown impurities (not previously detected in the same product prepared by state-of-the-art methods) should be below 0.10%, or completely qualified about structure and toxicity [5]. The impurities (organic related substances) may be unreacted starting materials, as well as intermediates and reagents used at different steps of the synthesis. In addition, degradation or by-products occurring during the process, in particular in harsh reaction conditions, might
Bioprocesses for the Synthesis of Nucleosides and Nucleotides
49
strongly influence the impurity profile of the final product. Also the presence of residual solvents should be carefully controlled. In fact, strict limits (ppm) are accepted for these contaminants due their toxicity [6]. Chemical methods [2, 3] used for the synthesis of modified nucleosides and nucleotides are usually based on a convergent approach via condensation of the carbohydrate precursor and the heterocyclic base, or employ natural nucleosides as starting compounds. In the chemical glycosylation, protection and deprotection steps and glycosyl activation are required for the control of the configuration at the anomeric center and the regioselective glycoside bond formation at one of the nucleophilic groups in the nucleobase. The regio-, chemo- and stereo-selectivity of the glycosylation reaction are ensured by the use of properly protected and activated sugars and bases that can be prepared by multistage processes. The classical chemical synthesis of pyrimidine ribonucleosides requires the preparation of the protected base (by silylation with hexamethyldisilazane) which is then reacted with 1β-acetyl-2,3,5-O-triacylribofuranoside (Figure 1). The nucleobase can be stereoselectively coupled to the sugar via the neighboring 2’-O-acyl group participation by the Vörbruggen glycosylation procedure. [2, 3] This synthesis must be reconsidered for the preparation of 2’deoxyribonucleosides, since the lack of a substituent in position C-2’ produces a 1:1 mixture of α/β anomers. For this reason, 2’-deoxyribonucleosides are usually synthesized via the ribocounterparts followed by 2’-deoxygenation. These syntheses are not very efficient for large scale productions. Each step requires chromatographic purification, the use of hazardous, expensive (i.e. TMSOTf as glycosylation promoter) or polluting reagents (i.e. Bu3SnH as reducing agent). The synthesis of purine nucleosides is even more complex and the strategy must be reconsidered when the target is the preparation of modified nucleosides. Generally speaking, selective modifications in the sugar moiety require several protection and deprotection steps. For example, by regioselective 3’-modification of thymidine it is possible to synthesize Zidovudine (AZT), [7] the first antiretroviral drug approved for the HIV treatment. [8] RO O RO
RO
OR1
Base
Glycosylation
Deprotection
O
OR
RO
HO
Base O
OR
HO
OH
R=Bz, R1 =Ac
Protection
HO
Base
Functionalization
OR
Deprotection
O RO
X HO
Base O Y
Figure 1. General pathway for the chemical synthesis of ribo- and 2’- or 5’-modified nucleosides.
5’-Monodeprotected nucleosides are used as intermediates to obtain 5’-mononucleotides [9] employed as food additives or antitumor prodrugs. [10-12] Similarly, nucleosides bearing
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50
a free hydroxyl group in C-5’ are used as precursors for the synthesis [13, 14] of (non natural) 5’-deoxynucleosides with antitumoral activity. [15, 16] In this case (Figure 2), the first step consists of the glycosylation between the peracetylated ribose and 5-fluorouracil or 5fluorocytosine, previously activated and protected by silylation. The protected 5fluorouridine or 5-fluorocytidine are then completely deprotected. Successively, both synthons are selectively protected at the C-2’ and C-3’ positions to allow the reduction of the free hydroxyl in C-5’ which occurs in two steps: treatment with triphenylphosphite methyl iodide and catalytic reduction. The final acid hydrolysis affords Doxifluridine and the precursor of Capecitabine in about 50% overall yield. For the synthesis of Capecitabine, the intermediate is treated with tert-butyldimethylsilyl chloride or simply by acetylation [14] to selectively protect the C-2’ and C-3’ positions. The resultant compound is subjected to N4-derivatization to give 2’,3’-bis-O-(tertbutyldimethylsilyl)-5’-deoxy-N4-(penthyloxycarbonyl)-5-fluorocytidine in high yield (92%) with a quite good impurity profile. Final deprotection of the sugar in mild conditions, to avoid the base deacylation, affords Capecitabine in 80% yield. As above described, the chemical synthesis of modified nucleosides and nucleotides is quite complex and may sensitively affect the quality of the target product. For this reason, simpler synthetic methods that do not produce isomers, by-products or other contaminants, including residual solvents, should be preferable. R
R
F
F
N N
AcO
OAc
O
R F N
OSi(CH3)3
N
N O
N
AcO O
R= OSi(CH3)3 R= NHSi(CH3)3
OAc OAc
O
HO O
OAc OAc
OH
OH
R= OH R= NH2
R= OH R= NH2 R
R
F
R
F N
N
F N
N HO
Triphenylphosphite methyl iodide
O O
O
O
N
N O O
a) H2, Pd/C; MeOH/TEA, O
H3C CH3
H3C CH3
R= OH R= NH2
R= OH R= NH2
Figure 2. (Continued on next page.)
O
I CH3
O
O
b) CF3COOH OH
OH
R= OH Doxifluridine R= NH2
Bioprocesses for the Synthesis of Nucleosides and Nucleotides O
O NH2
O
HN
F N
F
O
OR
N
O N
CH3
O
HN F
N N
51
CH3
O
N CH3
O
O
O
OR OR
OR
OH
OH
Capecitabine
Figure 2. Synthesis of 5’-deoxynucleosides used as antitumour agents.
In this context, enzymatic syntheses have been shown to be an advantageous tool, [17] also in combination with chemical methods. The use of enzymes as catalysts in pharmaceutical chemistry is quite new, but it is becoming popular also because of the recent attention for “sustainable chemistry”. In fact, enzymes efficiency carries out faster processes, less waste and a reduced use of organic solvents. Besides, the inherent enzymes’ selectivity as well as their ability to work in mild conditions can favour the obtainment of pure products by avoiding tedious synthetic and purification steps in comparison with the classical chemical approach. Thus, enzymatic processes can be very competitive in terms of costs and technology, allowing high quality products to be obtained. The availability of enzymatic “libraries”, the efficiency of molecular cloning and protein expression platforms, and all the technologies that can improve an enzyme’s selectivity, specificity and stability, are contributing to the diffusion of biocatalysis in the manufacturing of Active Pharmaceutical Ingredients (APIs). [18] Biotransformations can be performed by using the biocatalyst in one of the following “forms”: (i) whole cells, (ii) cell free extracts, (iii) isolated and purified enzymes, (iv) immobilized enzymes. Unlike conventional synthetic routes which can be normally controlled by means of reproducible chemical and physical techniques, biological processes, in particular when whole cells are used, are likewise variable. Thus, also in this case, full adherence to regulatory requirements for biological processes is mandatory for the efficient process control. The use of entire microorganisms pose many problems for the purification of the final product because of the co-occurance of reactions catalyzed by other enzymes as well as possible residues of the microorganism that can reduce the yield and the purity of the product. [19] Moreover, intra- and extra-cellular diffusion problems have to be considered. Alternatively, extracted and purified enzymes are conveniently employed to carry out reactions at laboratory scale. Many of the nucleosides and nucleotides bioconversions are performed by using free enzymes. Purified enzymes sensitively reduce some of the above mentioned drawbacks but, the instability of the protein in non physiological conditions (temperature, pH, ionic strength, cosolvents), the difficult recovery of the catalyst from the reaction medium and the high production costs, may represent important limitations. Besides, the enzymes in solution can be found as contaminants in the final product and this event is
52
Marco Terreni, Daniela Ubiali, Teodora Bavaro, et al.
not acceptable. For regulatory approval, bacterial contaminants are clearly defined (including residual proteins) and must be strictly controlled (especially for injectable dosage form). [19] Ultrafiltration could be necessary to ensure purification of the product from residual protein, in particular for water soluble products such as nucleosides and nucleotides. In this frame, the use of immobilized enzymes may be advantageous for developing industrial process. Immobilized enzymes are chemically bonded, or absorbed, to carriers (solid supports), or are entrapped in soluble form in devices, such as microcapsules or membranes, that are impermeable to the enzyme, ensuring a continuous exchange of substrate or product. These enzyme preparations can be separated from the reaction medium by simple filtration and are thus available for the re-use. On the other hand, it is possible that small changes in the exact shape of the active centre alter (with possible enhancement of) their catalytic properties. In fact, depending on the immobilization technique, the properties of the biocatalyst such as stability, selectivity, Km-value, etc. may be significantly affected. [20] Properly designed immobilization on solid supports enables the re-use of the catalyst and may increase the stability of many enzymes in a wide range of experimental conditions, including harsh conditions often required for the solubilization of substrates and products at the high concentrations necessary in preparative or industrial processes. Covalent immobilization may ensure a complete separation of the catalyst from the reaction medium by a simple filtration avoiding contamination of the solution containing the reaction product. On the contrary, enzymes only adsorbed on the matrix (by ionic or hydrophobic adsorption) can be released from the matrix, expecially in drastic conditions of temperature, presence of co-solvents and pH. Indeed, the proper design of a tailor-made immobilization plays a pivotal role to take full advantage from this technology. With a view to preparative (industrial) applications, it is mandatory to simplify the scale-up process; the availability of an immobilized and stabilized enzyme meets this need both for the easy handling of such a catalyst and its operational stability for repeated use. [21 a-c] This stability problem reaches a special relevance when using complex enzymes, as multimeric proteins that must keep their quaternary structure in order to preserve their catalytic activity. In fact, these enzymes are mainly inactivated by subunit dissociation, and this phenomenon may be enhanced under certain experimental conditions such as extreme pH values, low ionic strength, high temperature, organic solvents. Thus, it may be possible that a particular condition, where the enzyme might exhibit the most adequate performances, coincides with conditions where that multimeric enzyme tends to dissociate, therefore becoming inactive. In this case, also after covalent immobilization, the risk of protein release from the solid matrix cannot be excluded because not all the subunits are directly bonded to the support surface (Figure 2). Thus, for multimeric enzymes, the immobilization techniques used for the preparation of the catalyst should be carefully designed to successful assist the development of bioconversions at preparative-large scale, ensuring that no protein is released in the reaction mixture and preserving the quality of the final compound. In this case, stabilization of multimeric enzymes by immobilization and post-immobilization cross-linking can be achieved. [21 d-e]
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53
Cross-linking
•Dissociation •Dissociationof ofthe thesubunits: subunits: •Low stability of the •Low stability of theenzyme enzyme •Residual •Residualprotein proteinin inthe thefinal finalproduct product
No protein release
Figure 3. Dissociation of covalently immobilized multimeric enzymes in drastic reaction conditions can be avoided by post-immobilization cross-linking.
In this review, we report different examples of biotransformations used for the synthesis of nucleosides and nucleotides. In particular, we have considered the use of glycosyltransferring enzymes (e.g. nucleoside phosphorylases, N-deoxyribosyltransferases) for the synthesis of nucleosides by enantioselective transfer of glycosyl residues to acceptor bases. Moreover, hydrolases (e.g. lipases, esterases) are described in carrying out regio-controlled manipulations on the carbohydrate moiety. Finally, the enzyme-mediated synthesis of nucleotides through regioselective phosphorylation of the parent nucleosides catalyzed by kinases and phosphatases is also discussed. This review will also focus on the importance of the enzyme selection and the design of the final biocatalyst thereof, particularly by tailormade immobilization of the protein on solid support, with the aim to have an active, stable and reusable biocatalyst for preparative purposes.
2. Glycosyl-Transferring Enzymes 2.1. Nucleoside Phosphorylases: A Brief Overview about Properties and Functions The nucleoside phosphorylases catalyze the cleavage of ribonucleosides and deoxyribonucleosides to the free base (B1) plus ribose 1-phosphate or deoxyribose 1phosphate, respectively (Figure 4). The presence of an acceptor nucleobase (B2) results in the formation of a new nucleoside (transglycosylation). The reversible phosphorolysis of purine and pyrimidine nucleosides is an important biochemical reaction in the salvage pathway which provides an alternative to the de novo purine and pyrimidine biosynthetic pathways and a source of sugars for energy and carbon recovery. The bases serve anabolic (reutilization in nucleotide synthesis) or catabolic (use as nitrogen sources) functions.
Marco Terreni, Daniela Ubiali, Teodora Bavaro, et al.
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B1
HO
B1 HO E, Pi
O OH R
O OH ROPO3
E, B2
B2
Pi HO O
2-
OH R
B1, B2: purine or pyrimidine E: nucleoside phosphorylase R=H, OH Pi: inorganic phosphate Figure 4.
Structural studies have revealed that all of the enzymes that catalyze the phosphorolysis of nucleosides have only two possible folds. [22] This discovery provides the basis for defining two families of nucleoside phosphorylases. The first family (nucleoside phosphorylase-I, NP-I) includes enzymes that share a common single-domain subunit, with either a trimeric or a hexameric quaternary structure. They accept a wide range of purine nucleosides and uridine. Despite differences in substrate specificity, amino acid sequence and quaternary structure, all members of this family share a characteristic subunit topology. The second family of nucleoside phosphorylases (nucleoside phosphorylase-II, NP-II) share a common two-domain subunit fold, a dimeric quaternary structure, and a significant level of sequence identity (>30%). They are specific for pyrimidine nucleosides. NP-II accept both thymidine and uridine substrates in lower organisms, but are specific for thymidine in mammals and other higher organisms. The deep understanding of the enzymes active site and their catalytic mechanism is important to guide the selection of the enzyme as a catalyst to be used in a synthetic bioprocess.
2.2. NP-I Family: Structures, Active Site and Catalytic Mechanism Purine nucleoside phosphorylase (PNP; EC 2.4.2.1) is the most studied member of the NP-I family. PNP has been isolated and studied from a wide variety of both mammalian and bacterial species. These studies have revealed two forms of PNP: i) trimeric PNP, that is specific for guanine and hypoxanthine (2’-deoxy)ribonucleosides; and ii) hexameric PNP that accepts adenine, guanine and hypoxanthine (2’-deoxy)ribonucleosides. In bacterial species only the hexameric form has been largely found, with the exception of Escherichia coli, Bacillus subtilis and Bacillus stearotermophilus that possess both a trimeric and a hexameric form. [22] Uridine phosphorylase (UP; EC 2.4.2.3) and 5’-deoxy-5’-methylthioadenosine phosphorylase (MTAP; EC 2.4.2.28) belong to the NP-I family, too. UP is specific for uridine nucleosides, but also accepts 2’-deoxypyrimidine nucleosides. UP does not cleave cytidine, which is not cleaved by any known nucleoside phosphorylase, probably due the lack of hydrogen-bond-donor interactions with O-4 position. According to Krenitsky and coworkers [23], who studied pyrimidine nucleoside phosphorylases (PyNPs) from a broad range of bacterial and mammalian species, there exist two distinct pyrimidine cleaving enzymes:
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55
UP and thymidine phosphorylase (TP; EC 2.4.2.4). The latter, specific for the 2’-deoxyribose moiety of thymidine, belongs to the NP-II family. Despite this discriminating feature between UP and TP (the high specificity of TP for the 2’-position of the ribose moiety), for both enzymes the 5-position of the pyrimidine ring appears less important in determining specificity. MTAP is specific for the purine analogue 5’-deoxy-5’-methylthioadenosine. X-ray crystallographic studies have now provided several crystal structures of the NP-I family members in both native and complex forms. [22] Despite species-dependent differences in the quaternary structures of the NP-I family members, the subunit fold is highly conserved. The main feature of the common subunit fold of the NP-I enzymes consists of a central beta-sheet that forms a distorted beta-barrel, surrounded by several alpha-helices. [22] The active site consists of adjacent phosphate- and nucleoside-binding pockets. These binding sites are formed by residues from the central beta-sheet and the interconnecting loops, and residues from the adjacent subunit. It is significant that the central beta-sheet structural motif, where the nucleoside and the phosphate bind at the C-terminal end of the strands, is structurally conserved among the structures of the NP-I family. This suggests the importance of this motif in catalyzing the phosphorolysis. Previous work by Pugmire and Ealick [24] has reported a detailed comparison of the active sites and substrate binding modes in trimeric and hexameric PNPs [22]. Trimeric and hexameric subfamilies show significant differences in the positions of active-site residues (Figure 5). Although nucleoside synthesis is favoured thermodynamically, phosphorolysis generally occurs in vivo due to the uptake of the free bases as reactants in subsequent reactions [22]. Several different mechanisms for the phosphorolysis reaction have been suggested for members of the NP-I family from different species. While the members of the NP-I family all bind the phosphate and a nucleoside (which then undergo the phosphorolysis), the modes of substrate binding show significant variation, and help to explain the differences in substrate specificity. There is a similarity among the active site residues of trimeric structures (bPNPs), while the hexameric structures (E. coli PNP and UP, whose active site is not well characterized) appear to utilize different residues for binding the phosphate ion. The schematic drawings of a representative active site of each NP-I subfamily are reported in Figure 4. A schematic diagram of the proposed catalytic mechanism in hPNP, which is representative of a likely mechanism for all members of the NP-I family, is shown in Figure 5. The beta-nucleoside binds in a high energy +anticlinal torsion angle of the glycosydic bond, with the ribose moiety in the uncommon C-4’ endo sugar pucker. This high-energy conformation produces steric strain, which enhances glycosidic cleavage. The glycosidic bond is further weakened by electrons flow from O-4’ to the purine ring. This results in an oxocarbenium ion that is stabilized by the negative charges of the phosphate ion. The phosphate ion binds on the alpha-side of the ribose ring and participates in a SN1 nucleophilic attack at the C-1’ position.
56
Marco Terreni, Daniela Ubiali, Teodora Bavaro, et al.
Modified from ref. 22. Figure 5. Schematic drawings of a representative active site of each NP-I subfamily. a) Trimeric PNP; b) MTAP; c) hexameric PNP; d) UP. Shown are the residues that are thought to be involved in substrate binding.
Bioprocesses for the Synthesis of Nucleosides and Nucleotides
H 2N
O
O
δ- N δ+ O
O
NH
N N
HO HO
57
OH O P O OH
-
Modified from ref. 22. Figure 6. Proposed catalytic mechanism for hPNP, representative of a likely mechanism for all NP-I family enzymes.
Kinetic studies of UP from different species have reported both a random-sequential mechanism [22] and an ordered-sequential mechanism in which phosphate is the first substrate and uracil is the first product to leave. Both mechanisms have been purposed also for MTAP.
2.3. NP-II Family: Structures, Active Site and Catalytic Mechanism Detailed structural information on the NP-II family was initially provided by the crystal structure of E. coli TP. [25] The deoxyribosyltransferase activity was first attributed to TP by Zimmerman and Seidenberg. [26] Later studies verified this activity, and also showed that the 2’-deoxyribose 1-phosphate and the pyrimidine base bind at two distinct locations in the enzyme. [27] It was further shown [28] that the deoxyribose 1-phosphate is a free intermediate, so that the transferase activity, involving two pyrimidine bases B1 and B2, is likely to take place via a two-step mechanism as reported in Figure 6. Kinetic studies of TP from several species generally agree on a sequential mechanism. Studies of TP from E. coli [28] and Lactobacillus casei [29] suggest an ordered sequential mechanism, with phosphate binding before thymidine, and thymine being released before deoxyribose 1-phosphate. These results are corroborated by evidence that phosphate or deoxyribose 1-phosphate stabilizes the enzyme, but thymidine or thymine does not. [30] TP functions as a dimer consisting of two identical subunits related to each other by a 2fold axis of symmetry and an S-shape. [31] Each subunit contains a large mixed α-helical and a β-sheet domain (α/β-domain) separated from a smaller α-helical domain (α-domain) by a large cleft (Figure 7).
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Marco Terreni, Daniela Ubiali, Teodora Bavaro, et al.
Source: http://arginine.chem.cornell.edu/Structures/TP.html. Figure 7. Ribbon drawing of the dimeric structure of E. coli TP in an open conformation.
Three loop regions connect these two domains. The active site of each subunit consists of a thymidine binding site in the α-domain and a phosphate binding site across the cleft in the α/β-domain. The large distance between the phosphate and the thymidine binding sites reported in E. coli TP suggested that rigid-body domain movement, where the loop regions connecting the two domains act as hinge regions, might close the cleft and enable catalysis to proceed. Evidence of the proposed rigid-body movement of the two domains has been reported for E. coli TP, in that it two domains have moved under the packing constraint of different space groups [24]. This movement of the two domains towards each other would decrease the distance between the phosphate and thymidine and enable the catalysis to take place. [24] Further evidence for domain movement and subsequent cleft closure was obtained with the crystal structure of PyNP from B. stearothermophilus which was found to be very similar to E. coli. [24] Based on structural studies, binding of phosphate in E. coli causes an initial rotation of the α/β-domain through the formation of a hydrogen bond between the sidechain of H119 and the backbone carbonyl oxygen of G208 (H116 and G205 in B. stearothermophilus). Binding of the pyrimidine causes an additional rotation, which produces the cleft closure in the enzyme active conformation. The possible mechanism of domain movement triggered by binding of substrates in B. stearothermophilus is reported in Figure 8.
Bioprocesses for the Synthesis of Nucleosides and Nucleotides
59
α/β T154 NH
α/β Q 153
L1
L2
L3
T154
L2
L1
NH
O H 2N
O L157
+ H 3N
O
D161
O
+
O
L15 7
O
α
K 187
L3
Thd H 3N
O D161
α
K187
Modified from ref. 22. Figure 8. Ribbon drawing of PyNP from B. stearothermophilus. The α/β domain is shown in red, the α domain in blu and in green are depicted the three loops that act as hinge regions. Phosphate and uracil are shown as sphere models to indicate the position of the active site. In the schematic representation, the pyrimidine substrate is indicated as a gray circle. The three hinge regions (green) are labelled L1L3. The proposed hydrogen bond formation that forms a β turn is indicated by the dashed line in the L2 region.
Binding of phosphate causes a partial domain closure. A further domain movement is triggered when pyrimidine binds in its binding site. The sidechain of K187 changes conformation so that the hydrogen bond with the pyrimidine ring can form. Prior to substrate binding, K187 is positioned to form a salt bridge with D161. Once this salt bridge is disrupted by substrate binding, D161 forms a hydrogen bond with Q153. The formation of this hydrogen bond coincides with the formation of a β turn in the L2 hinge, where a hydrogen bond forms between the carbonyl oxygen of L157 and the backbone nitrogen of T154. The phosphate-binding pocket is located between two β-strands near the C-terminal end of the central β-sheet of the α/β-domain (Figure 8). Residues that bind phosphate come from these two strands as well as from a nearby loop. The pyrimidine-binding pocket is formed
Marco Terreni, Daniela Ubiali, Teodora Bavaro, et al.
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from residues in the two helices on the face of the α-domain facing the cleft. The primary interactions include binding of the two carbonyl groups of the pyrimidine ring by positively charged arginine and lysine residues, as well as a herringbone base-stacking interaction between the pyrimidine base and a tyrosine residue. The crystal structure of TP from E. coli and PyNP from B. stearothermophilus complexed with a phosphate ion and pyrimidine base along with computational studies of the nucleoside modelled into the active site have identified the residues involved in binding the substrates (Figure 9) [24, 25]. These residues are highly conserved among all the enzyme of the NP-II family. Comparison of the residues that bind the sugar moiety in E. coli and human TP (that are specific for 2’-deoxyribose) and in PyNP (that is able to accept both ribose and 2’deoxyribose), suggests that the substrate specificity is due to a methionine that, replacing K108, may create a different hydrogen-bonding scheme with the phosphate oxygen that binds the 2’-hydroxyl of the ribose moiety (Figure 6). Moreover, even if the structure of NP-I and NP-II enzymes are quite different, the overall orientation of the nucleoside and the phosphate ion in the active site in the closed enzyme conformation are similar [24)]. This suggests that the catalytic mechanism is very similar to those proposed for the NP-I family. A proposed mechanism for NP-II family is shown in Figure 10. A rg 1 6 8 T yr1 6 5
NH
H 2N
NH 2 H O
O
S e r1 8 3
NH HO N HO
O H 3N
+
O OH
S e r8 3
O
NH NH
O-
P O-
L ys 8 1
OH
3
+
O-
+
OH
HN
O H
N H is 8 2 S e r1 1 0
H 3N
T h r1 2 0 CH
3
L ys 1 0 8 (M e t)
Modified from ref. 22. Figure 9. Active-site residues in the PyNP structure of B. stearothermophilus in a closed-cleft conformation. Hydrogen bonds are shown as dashed lines.
Bioprocesses for the Synthesis of Nucleosides and Nucleotides
61
Arg171 H2N
NH2 δO
H3C
Ser186
NH Oδ
-
δ+ N O
HO
OH
Lys190 Ser86
OH OH H OO O- P OH O O
Lys84
O H
Thr123
OH Lys191 Asp83
Figure 10. Schematic drawing of the proposed model for catalytically active thymidine phosphorylase, based on the simulated closed-cleft model. Dotted lines indicate possible hydrogen bonds.
2.4. Other Glycosyl-Transferring Enzymes: N-Deoxyribosyltransferases (DRTases) N-Deoxyribosyltransferases (DRTases) (EC 2.4.2.6), also called trans-Ndeoxyribosylases, catalyze the direct transfer reaction of a 2’-deoxyribosyl moiety from a donor deoxynucleoside to an acceptor nucleobase (Figure 11). [32, 33] B1
HO O
B1 HO E
OH
O OH
B2
E HO B2 E
O OH
B1, B2: purine or pyrimidine E: N-deoxyribosyltransferase Figure 11.
N-Deoxyribosyltransferases have been found mostly in lactic acid bacteria, such as Lactobacillus and Lactococcus species [34, 35], although they are also found in the protozoan Crithidia lucilae. [36] Since DRTases are distributed in microorganisms that lack purine and pyrimidine nucleoside phosphorylases, which are used for nucleosides recycling in most organisms, it is thought that their physiological role is to scavenge exogenous 2’deoxynucleosides in the nucleoside salvage pathway for their DNA synthesis. DRTases have been divided into two classes on the basis of their substrate specificity. [37]
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DRTases I class, also called purine deoxyribosyltransferase (PTD), catalyze the transfer exclusively between purine bases with a specific preference order of dI> dA> dG concerning the deoxyribonucleoside donor. DRTase II class, also called nucleoside deoxyribosyltransferase (NTD), has substrate specificity to both purine and pyrimidine bases as donor and/or acceptor moiety, with a preference for deoxypyrimidines as sugar donor. Among N-deoxyribosyltransferases, NTD from Lactobacillus leichmanni and PTD from Lactobacillus helveticus are well characterized concerning their structure and reaction mechanism. The structure of NTD from L. leichmanni was determined by Armstrong et al. [38] The enzyme resulted to be a hexamer in its native state, composed of identical 18 KDa subunits, packed as a trimer of dimers. Anand et al. [39] resolved the structure of PTD from L. helveticus: PTD is also a hexamer. Catalysis proceeds via a ping-pong bi-bi mechanism (at least one product is released from the enzyme before all of the substrates have bound) with formation of a deoxyribosyl-enzyme intermediate. The substrate binding geometry and the key residues involved in catalysis are very similar in both the enzymes. In the active site three acidic residues are present which are implicated in substrate binding and catalysis, the hydrophilic core is shielded by a shell of hydrophobic residues [39]. In the formation of the covalent intermediate, one amino acid would function as a general acid/base and the second carboxylate (glutamyl or aspartyl residue) would serve as the attacking nucleophile. In particular, in the case of PTD, the nucleoside is oriented for catalysis by several hydrogen bonds. The cleavage of the N9-C1’ bond probably occurs via an oxocarbenium transition state in which negative charge accumulates on the purine ring and a positive charge accumulates on the C4’-O4’ bond of the sugar (Figure 12). T yr
O -O
H
A sp
NH H N
HO
N
δ-
O
N H
O δ +O
O-
N H O
Asp
OH O
-O
-
G lu
Figure 12. Proposed catalytic mechanism for DRT.
The N7 is protonated by a carboxylate group. The delocalization of the negative charge in the base ring may be further assisted by protonation of the N3 by aspartic residue. A particular feature of this enzyme is its ability to transfer the deoxyribose moiety not only to
Bioprocesses for the Synthesis of Nucleosides and Nucleotides
63
the usual N9 position but also to the N7 or N3 atom of the purine base. [39] This is possible because the plane of the purine base is the same in all three orientations. Both PTD and NTD are characterized by high stereospecificity of the glycosyl transfer, leading to the retention of the anomeric configuration (only the β anomer of the nucleoside is produced). DRTases are specific for 2’-deoxynucleosides; it has been hypothesized that the presence of a hydroxyl group at the C2’ position may hamper catalysis by forming a hydrogen bond with both the oxygen atoms of the active site nucleophile (glutamyl or aspartyl residue), but also by breaking a key hydrogen bonding interaction that results in the destabilization of the transition state. 2’,3’ Dideoxynucleosides are also competent sugar donors, but the transfer reactions are slower because the interaction of the 3’-OH with the active site residues, although not necessary, may help position of the sugar in the optimal orientation for catalysis [38].
2.5. Synthesis of Nucleosides by Enzyme-Catalyzed Transglycosylation or Glycosylation Enzymatic syntheses based on glycosyl-transferring enzymes have been shown to be an advantageous alternative to chemical methods due to the high selectivity of the enzymes, the mild reaction conditions and the overall simplicity of the approach. [17] NPs catalyze the reversible phosphorolysis of a nucleoside leading to the intermediate sugar-1’-phosphate with the release of the nucleobase. The addition of an acceptor base to the reaction mixture results in the formation of a new nucleoside (transglycosylation). [17, 40, 41] Coupled NPs system with a different specificity can be used at a preparative scale to synthesize a variety of base- and sugar-modified nucleosides (see Figure 4). Since the 60’s, when the enzymatic equilibrium-transfer reaction of the sugar moiety of one nucleoside to the heterocyclic base was first discovered by Krenitsky [42], it was established that this equilibrium reaction is catalyzed by PNP in the case of purine nucleosides and purine bases, and by UP or TP in the case of pyrimidine nucleosides and pyrimidines. The substrate specificities and synthetic applications of NPs toward modified purine, pyrimidine and ribose derivatives have been studied intensively. Several nucleoside N-transfer reactions via those phosphate esters of sugar were transribosylation, trans-2’deoxy-ribosylation, transarabinosylation, trans-2’,3’-dideoxyribosylation and trans-alpha-L2’,3’-dideoxyribosylation by NPs [43]. N-deoxyribosyltransferases provide an alternative to the nucleoside phosphorylases. The two transferases types (type I and type II) [37], which have different specificity, molecular weights and thermal stabilities [44], offer a different route for the synthesis of nucleoside analogues (see Figure 11). The use of enzymes or the whole cells as biocatalysts has resulted in a dramatic improvement of the key–step of the nucleosides synthesis, being the glycosylation reaction. This is due to the fulfillment of the stereo- and regio-selectivity occurring in the enzymecatalyzed formation of the C-N bond. The chemical-enzymatic technology has been shown to sensitively improve the cost-effectiveness ratios versus chemical processes. In this context,
64
Marco Terreni, Daniela Ubiali, Teodora Bavaro, et al.
the design of the optimal biocatalyst for efficiency, operational stability and costs plays a crucial role. Considerable progress in the preparation of nucleoside analogues has been achieved by combination of chemical methods and biochemical transformations [45].
Nucleoside Phosphorylases (NPs) The enzymatic preparation of nucleosides by using uridine phosphorylase (UP) or thymidine phosphorylase (TP) and purine nucleoside phosphorylase (PNP) has been extensively described in literature as well as claimed in some patents. [46] Besides, a constant interest is being devoted to the search of new microorganisms endowed with a wide substrate specificity and, therefore, capable of synthesizing modified nucleosides from available precursors. For instance, the extensive microbial screening recently performed by Trelles et al. has provided interesting results at analytical scale [47]. With the aim to somehow classify the most relevant examples of enzymatic glycosylation, we have considered the following criteria: the advantage of the bioprocess versus the standard procedure and the feasibility of the bioprocess at a preparative (industrial) scale, which also includes the evaluation of the biocatalyst performance. For example, in WO 00/39307 [46a], natural nucleosides are synthesized by recombinant E. coli cells which overexpress the enzymes UP and PNP. This method avoids the enzyme extraction and purification steps and also allows the sufficiently easy recovery of the cells which can thus be used for successive production cycles. However, the number of cycles is limited by the high working temperatures (60°C) which are used to ensure the acceptable solubilization of the substrates and also to deactivate adenosine deaminase (ADA), to avoid unwanted side-reactions. The use of cell pastes is also generally constrained by some limitations represented by intra- and extra-cellular diffusion problems and by the use of large quantities of cell suspension which reduces the volumetric yields of the final product. Finally, a further drawback is due to the possible presence, in the final product, of contaminants such as proteins of the microorganism, side-reactions products or toxins. For these reasons, the use of isolated enzymes is generally preferred even though, in this case, other drawbacks, such as the poor stability of the free enzyme in non physiological conditions, have to be considered. Barai et al. [48] recently reported the synthesis of Cladribine (4, 2-chloro-2’deoxyadenosine) and Ribavirin (5, 1-(beta-D-ribofuranosyl)-1,2,4-triazole-3-carboxamide) by transglycosylation starting from 2’-deoxyguanosine (1, 2’-dG) and guanosine (2, Gua), respectively, as sugar donor (Figure 5). The reactions were carried out by using E. coli BMT4D/1A cells that possess high UP and PNP activities. The cells were cross-linked by glutaraldehyde in order to have a stable biocatalyst for repeated use: this biocatalyst could be used for 5-10 cycles at 60-65°C. Cladribine(4) was isolated in 81% yield, Ribavirin (6) in 6770% yield. Hennen and Wong [49] described the successful use of isolated PNP in solution for the synthesis of adenosine (6), Ribavirine (5) and other modified purine nucleosides by transglycosylation starting from 7-methyl guanosine (3) (or 7-methyl inosine, here not reported) as donor nucleosides (Figure 13).
Bioprocesses for the Synthesis of Nucleosides and Nucleotides
65
H2N H2N N
N
X N N
N N H A, Z=H
Y
-O
Z
N
+ Z
N
N
NH
+
Y
NH2
N
N H
HO
OH R
2-Cl A, Z=Cl NH2
O
X N
O
N
HO
N
N
4, R=H Z=Cl
(Cladribine)
X=H
6, R=OH Z=H
(Ade)
X=CH3
+ O H2NOC
OH R
N
1, R=H X=H Y= -
(2'-dG)
2, R=OH X=H Y= -
(Gua)
3, R=OH X=CH3 Y=HI (7-M Gua)
N
H2NOC
N N N H
TCA
N HO
+
O
X N N H
O NH N
+
Y
NH2
OH R 5, R=OH
(Ribavirine)
X=CH3, H
Figure 13.
Pyrimidine-pyrimidine transglycosylation has been recently reported by using the native TP from E. coli. [50] An interesting investigation about the substrate specificity of this enzyme was performed with the aim to assess if the process was a feasible synthetic method for development of unnatural nucleosides. Figure 14 depicts all the base analogues which were converted to the corresponding nucleosides (10-19) in good yield (47-95%) as well as the effect of the hydroxyl group of the ribosyl moiety on the catalytic reaction by using 5fluorouracil as standard acceptor base (20-21). As previously reported (Paragraph 2.1), 5-substituted uracil compounds are generally well tolerated by TP, with the exception of 5-nitrouracil. Also 6-aza compounds, in which the carbon was replaced by a nitrogen atom (azauracil, azathymine), were not transformed as well as other C-6 substitutions like methyl or keto groups (5,6-dimethyluracil, 5,6dihydrouracil, barbituric acid) were not accepted (data non shown). 2-S-Substituted uracil (2thiouracil and 2-thiothymine) were instead recognized as substrates by TP to give the counterpart nucleosides 18 and 19. The investigation of the hydroxyl group effect of the sugar moiety concluded that the 5’OH is not required to recognize the substrate for TP. In fact, the transglycosylation from 8 (5’-deoxythymidine, 5’-dThd) and 5-fluorouracil to 20 occured in 85% yield in 3 hours. Similarly, 5-fluorouridine (21) was obtained in 82% yield from 5-methyluridine (5-MUrd, 9), showing that the 2’-alpha-OH does not influence the recognition of the ribosyl substrate by TP. Only the 3’-OH seems to affect the reactivity of this enzyme: when 2’,3’dideoxythymidine is used as sugar donor, no conversion is observed (data not reported). Pyrimidine nucleosides were largely used as sugar donor also for the preparation of purine nucleosides. A significant example is reported in Figure 15 for compounds 1 and 2732 [41, 51-54].
Marco Terreni, Daniela Ubiali, Teodora Bavaro, et al.
66
O CH3
HN O HN O R1
O
CH3
Y N H
Y N
+
O
X
HN Z
X
HN Z
N
R1 O
N H
OH R
OH R uracil
10, X=H Y=O Z=O (2'-dU)
5-fluorouracil
11, X=F Y=O Z=O (Floxuridine)
5-chlorouracil
12, X=Cl Y=O Z=O
5-bromouracil
13, X=Br Y=O Z=O (Broxuridine)
5-iodouracil
14, X=I Y=O Z=O (Idoxuridine)
5-ethyluracil
15, X=Et Y=O Z=O
5-aminouracil
16, X=NH2 Y=O Z=O
5-trifluoromethyluracil
17, X=CF3 Y=O Z=O (Trifluridine)
2-thiouracil
18, X=H Y=O Z=S
2-thiothymine
19, X=CH3 Y=O Z=S
5-fluorouracil
20, X=F Y=O Z=O
9, R=OH R1=OH (5-MUrd) 5-fluorouracil
21, X=F Y=O Z=O
7, R=H R1=OH (Thd)
8, R=H R1=H (5'-dThd)
Figure 14. Krenitsky et al. [41] combined UP or TP and PNP from E. coli in solution to synthesize the 2’-deoxyribonucleoside of 6-(dimethylamino)purine (25) and the ribonucleoside of 2amino-6-chloropurine (26) in yields of 81 and 76%, respectively, starting from thymidine (7) and uridine (22) (Figure 15). Nucleosides 1 and 27-32 were enzymatically synthesized by the same general pathway (pyrimidine nucleoside-purine base transglycosylation) but through the use of immobilized biocatalysts. As previously described, in fact, the immobilization of enzymes on solid support has many advantages: it may facilitate the protein recovery, it avoids the protein release in the reaction medium and it may enhance the enzyme stability. Stability is crucial, particularly for multimeric enzymes that, in certain reaction conditions (pH, organic solvents, temperature) can easily undergo subunit dissociation. The alteration of the quaternary structure results in the loss of enzyme catalytic activity. In spite of many reports have been published so far about enzyme immobilization [20, 21 a-c] and multimeric enzyme immobilization [21 d-e], only few examples have been described about the efficient
Bioprocesses for the Synthesis of Nucleosides and Nucleotides
67
immobilization of glycosyl-transferring enzymes and their use thereof in preparative bioprocesses also of industrial relevance. O X
HN O O
O
N
HO
W
X=H, CH3
X
HN
N H N
W N +
O R1
N H
N
N N
HO Z
N N
Z
O R1 OH R
OH R 7, R=R1=H X=CH3 (Thd)
6-(dimethylamino)purine
25, R=R1=H W=N(CH3)2 Z=H
22, R=OH R1=H X=H (Urd)
2-amino-6-chlopurine
26, R=OH R1=H W=Cl Z=NH2
23, R=H R1=OH X=H (ara-U)
6-methoxyguanine
27, R=H R1=OH W=OCH3 Z=NH2
adenine
28, R=H R1=OH W=NH2 Z=H (ara-A)
2,6-diamino-purine
29, R=H R1=OH W=NH2Z=NH2 (ara-DAMP)
2-fluoroadenine
30, R=H R1=OH W=NH2Z=F (Fludarabine)
hypoxanthine
31, R=R1=H W=OH Z=H (2'-dI)
guanine
1, R=R1=H W=OH Z=NH2 (2'-dG)
adenine
32, R=R1=H W=NH2 Z=H (2'-dA)
24, R=R1=H X=H (2'-dU)
Figure 15.
Problems of instability of the enzymes at high temperature were solved by Mahmoudian [51] who described the preparation of the 6-methoxy precursor of 9-beta-D-arabinofuranosyl guanine (27) starting from 6-methoxy guanine and arabinosyluracil (23, ara-U, Figure 7). UP and PNP from E. coli were not stable, as crude lysates, at the required reaction temperature (55°C); the enzymes were thus stabilized by direct co-immobilization onto an ion-exchange support (DEAE-52). This co-immobilized strategy allowed to set the basis for a scaleable process. No mention was made by the Author about the effect of pH on the enzymes’ stability, prior and after immobilization. However, the reaction was run at pH 7.4. Similarly, Hori et al. [55] immobilized a PyNP and two kinds of PNP (PNPI and PNPII) from B. stearothermophilus JTS 859 on an anionic exchange resin (DEAE-Toyopearl). These biocatalysts were used to prepare 5-methyluridine (9, 5-MUrd) starting from inosine (33) and thymine through the transglycosylation pattern purine nucleoside-pyrimidine base (Figure 16).
Marco Terreni, Daniela Ubiali, Teodora Bavaro, et al.
68
HO N HO N N HO O
N H
N N
N
CH3
HN O
N H
CH3
HN
Hpx
O
+
N
O
O
N
HO O OH OH
OH OH 33, (Ino)
thymine
9, (5-MUrd)
Figure 16.
Due to the thermophilic nature of the selected microorganism, the reactions could be performed at 60-70°C even with the soluble enzymes. However, the half-life of the immobilized enzymes was higher than that of the not-immobilized enzymes allowing the reuse of the biocatalysts. Interestingly, thermal stability was not accompanied by a similar trend about the pH resistance. Whereas the optimal temperature of the immobilized enzymes was shifted to a higher value (probably due to a stabilization induced by the immobilization), the optimal pH was very narrow in comparison with the crude enzymes. The Authors attributed this effect to the positive electrostatic field by the anion exchange resin on the local pH in the domain of the immobilized enzyme, in agreement with previous works [56]. A successful example of co-immobilization of UP and PNP by covalent linkages through epoxy-activated solid supports has been recently reported. [52] Epoxy-activated resins are ideal supports for protein immobilization due to the chemical stability of oxirane groups functionalized on the matrixes and their ability to react with amine and thiol groups on the protein surface to give stable C-N and C-S bonds. Accordingly, successful immobilization requires a preliminary physical interaction between the protein and the solid support through, for example, hydrophobic adsorption which can be promoted by the use of hydrophobic resins in the presence of high salt concentrations. [57] Covalent immobilization is more stable than the ionic interaction between the enzyme and the resin [51, 55] and thus can avoid risk of contamination from adventitious agents (protein release). By using co-immobilized biocatalysts, beta-D-arabinofuranosyl-adenine (28, ara-A, Vidarabine), beta-D-arabinofuranosyl-2,6-diamino-purine (29, ara-DAMP), beta-Darabinofuranosyl-2-fluoroadenine-des-phosphate (30, Fludarabine-des-phosphate), 2’deoxyinosine (31, 2’-dI), 2’-deoxyguanosine (1, 2’-dG), 2’-deoxyadenosine, (32, 2’-dA), were synthesized in a 70-85% yield and >99% purity for reaction times of 2-6 hours (Figure 7). These biocatalysts were characterized by a significant heat stability (the transglycosylations were routinely performed at 60-70°C), fully retained their enzymatic activity in the presence of organic solvents (DMSO, DMF, THF, PEG) and showed high reproducibility in consecutive bioconversions, all industrial-feasible conditions. Recently, our research group developed a novel immobilization technique for the immobilization of purified UP from Bacillus subtilis [53] and its use, together with covalently
Bioprocesses for the Synthesis of Nucleosides and Nucleotides
69
immobilized PNP, in the synthesis of 2’-deoxyinosine (31, Figure 15) and 2’-deoxyguanosine (1, Figure 7) by transglycosylation starting from 2’-deoxyuridine (24). [54] Aim of this research was to efficiently immobilize and stabilize the multimeric UP and PNPI from B. subtilis, in order to have stable biocatalysts under harsh (industrially-feasible) reaction conditions (Figure 15). The considered enzymes PnpI/PupG (PNP), with specificity for riboand deoxyribo- guanosine and inosine, and Up/Pdp (UP), with specificity for uridine, thymidine and deoxyuridine, from the purine and pyrimidine salvage pathway of B. subtilis were overexpressed in E. coli, purified, and successfully immobilized on solid support [53, 54], after an extensive study about the stabilization of the quaternary structure. To stabilize the multimeric UP, different commercially available supports, such as ionic agarose, epoxy resins or glyoxyl-agarose (derivatized with aldehyde groups), were used in order to study the effect of different matrixes and immobilization mechanisms. The simplest strategy was ionic adsorption on agarose DEAE, in agreement with the previous cited reports. [51, 55] However, the resultant biocatalyst was particularly unstable due to the weak interaction with the support bearing the ionic groups directly bond to the surface. This drawback was settled by using an epoxy resin (Sepabeads) coated with polycationic polymers (polyethylenimine), allowing the enzyme to be fully covered by the polymer. [57, 58] To further increase the stability of aminated support-enzyme complex and prevent enzyme desorption, the enzyme derivative was treated with polyaldehyde macromolecules (20% oxidized dextran) that, reacting both with the free amino groups of the enzyme and polyethylenimine, afforded a covalent multipoint cross-linking between the protein subunits and the support. The activity and the stability of this biocatalyst was very high compared with the non cross-linked derivative (Figure 17).
120
Activity %
100 80 DEAE-agarose
60
Sepabeads-PEI-dx
40
Free UP
20 0
0
5
10
15
Time (hours)
Figure 17. Stability of UP: pH 10 and 45°C.
20
25
Marco Terreni, Daniela Ubiali, Teodora Bavaro, et al.
70
The stabilized UP preparation retained up to 70% of the total activity at pH 10 and 45°C whereas the immobilized UP by ionic adsorption (DEAE agarose) and the non immobilized enzyme were promptly inactivated under the same conditions. The yields of 2’-dI (31) and 2’dG (1) were 85 and 95%, respectively. The first, obvious approach to enhance the bioconversion efficiency is not only to modulate the substrate concentrations ratio, but also to somehow remove the product(s). In US 006017736A [59], an interesting method for preparing purine nucleosides by enzymatic transglycosylation is claimed. The reaction involves a pyrimidine nucleoside as a sugar donor and a purine base as acceptor. PyNP and PNP are used, indeed, as catalysts. Interestingly, the pyrimidine base formed by the base exchange reaction is converted by uracil thymine dehydrogenase (E.C. 1.2.99.1) or dihydrouracil dehydrogenase (E.C. 1.3.1.1) into a compound which is not a substrate of the used enzymes. As a result, the equilibrium of the reaction is shifted towards the purine nucleoside which is formed in high yields. Certain enzymatic activities, not “directly” involved in the transglycosylation, may be exploited to affect the equilibrium shift as described above or also to catalyze the conversion of a “pro-substrate”. In the chemo-enzymatic synthesis of ara-G (35, Figure 18) [48, 60], a concerted pathway of three biochemical reactions is involved: cytidine deaminase (E.C. 3.5.4.5, CD), UP and PNP. Ara-C (34) was used as a donor of the beta-D-arabinofuranose moiety and guanine as an acceptor (Figure 18). O HN O
O
N
HO
N +
N
N
N H
O OH
N
HO NH 2
UP/PNP
O OH OH
OH 23, (ara-U)
35, (ara-G)
guanine
CD
PNP NH 2
HO
N O
N N
HO O OH OH 34, (ara-C)
Figure 18.
N
HO
HN O
HO
N H
N HO
N N
O OH
R
1, R=H 2'-dG 2, R=OH Gua
NH 2
N N
NH 2
Bioprocesses for the Synthesis of Nucleosides and Nucleotides
71
Since ara-C (34) is not a substrate for UP, in order to transfer the beta-D-arabinofuranose from ara-C (34) to the purine base, the former is first deaminated to 9-(beta-Darabinofuranosil)uracil (ara-U, 23). In fact, the E. coli BM-11 cells were selected due to their high content of CD activity along with UP and PNP. Ara-U (23) is therefore subjected to phosphorolysis to give arabinosylfuranose-alpha-1-phosphate. This intermediate was stereoand regio-selectively coupled with guanine to afford ara-G (35). Guanine was formed in situ under the action of PNP on 2’-dG (1) or Gua (2), employed as soluble source of this purine base. A similar strategy was exploited for the preparative-scale synthesis of 9-(3-deoxy-betaD-erythro-pentofuranosyl)-2,6-diaminopurine (38), important precursor of related nucleosides and 2’,5’-phosphodiester linkage oligonucleotides. The reaction was performed from 3’-deoxycytidine (36, 3’-dCyt) and 2,6-diaminopurine by using E. coli BMT-4D/1A cells at pH 7.0 and 52°C (Figure 19) [61]. O
NH2 N O
HN
H2 N N
N
a
N
+
HO
N H
O
N
O N
O O
NH2
HO
HN
HO
O
N H
+
O OPO32OH
OH
OH 36 (3'-dCyt)
37 (3'-dU)
2,6-diaminopurine
O
H2N N
N N
NH2
N
NH
N
b
N
N
NH2
HO
HO
O
O OH
OH 39 (3'-dG)
c
38
H2N d
N N
H2N N N HO
HO
N N
N N H
O
O F OH 40 (3'-disoG)
O 41
OH
Figure 19.
Reaction conditions: a: K phosphate buffer pH 7.0, 52°C, 26 h, yield 64%; b: ADA, r.t., 16 h (a+b, yield 67%); c: NaNO2/AcOH, 50°C, 6 min. yield 71%; d: HF/HBF4/H2O/THF NaNO2, -10,-12°C, 1h, yield 43%.
72
Marco Terreni, Daniela Ubiali, Teodora Bavaro, et al.
A preliminary deamination of 3’-dCyt (36) (due to the presence of CD activity in the intact cells) led to the formation of 3’-dU (37) which underwent to phosphorolysis affording the sugar for the transglycosylation with 2,6-diaminopurine. The yield of the target nucleoside, 2,6-diamino-9-(3-deoxy-beta-D-erythro-pentofuranosyl)purine (38), after purification by column chromatography, was 64%. The Authors stressed the remarkable advantages of this method over the chemical glycosylation of the 2,6-diaminopurine derivatives, which lacks regio- and stereo-selectivity. It is noteworthy that 2,6-diamino-9-(3deoxy-beta-D-erythro-pentofuranosyl)purine (38) is a useful synthon for the preparation of high-value related nucleosides: 3’-deoxyguanosine (39, 3’-dG, 85% yield by cells containing ADA activity), 3’-deoxyisoguanosine (40, 3’-disoG, 71% yield by NaNO2 treatment in AcOH/H2O) and 9-(3-deoxy-beta-D-erythro-pentofuranosyl)-2-fluoroadenine (41, 43% by Schiemann reaction). This example emphasizes how, although several methods of stereoselective glycosylation have been developed so far, glycosylation with a guanine base is still an ardous task, especially for application to scaleable preparations. As mentioned in Figure 19, the synthesis of guanine derivatives can be performed by using adenosine deaminase (ADA). ADA (E.C. 3.5.4.4) is another enzyme that has been considered for the nucleosides synthesis, since it catalyzes the rapid and irreversible deamination of adenine nucleosides as well as other C6 purine analogues to the corresponding hypoxanthine ones. [62] A review about the role of ADA in nucleoside synthesis has been recently published. [63] An interesting example of a preparative bioprocess through immobilized adenosine deaminase has been reported. [64] Commercial ADA (from calf intestinal mucosa and Aspergillus sp.) was immobilized on Eupergit C for the synthesis of Carbovir starting from the carbocyclic nucleoside aristeromycin. The enzyme was reused up 10 cycles without any significant loss of activity to give Carbovir in >90% yield. Enzymatic reactions with guanine derivatives, often coupled with few chemical steps, are very promising. The importance of this strategy is supported by few examples where a chemo-enzymatic process was developed to replace an existing and less efficient chemical route. This is the case of 2’,3’-dideoxy-3’-fluoro-beta-D-guanosine (44) [65], which has been showed to have a prominent antiviral activity. [66] The enzymatic glycosylation (Figure 20) was directly performed by reacting guanine and 2,3-dideoxy-3-fluoro-alpha-D-ribose 1phosphate (42), stereoselectively synthesized as described in the same work. The use of bacterial PNP in aqueous solution afforded the modified nucleoside in high conversion. The product was isolated by crystallization directly from the reaction mixture in pure form (final two-steps yield: 63%). The efficiency of this strategy relies on the fact that no isomers of the target compound were formed, thus providing an alternative synthetic route for the preparation of unnatural nucleosides. In fact, the same Authors [67] successfully applied the methodology above described to the synthesis of all four natural 2’deoxynucleosides. The key-compound, being the 2-deoxyribose 1-phosphate (43), was prepared with high stereoselectivity by the novel method [68] and then reacted with each natural nucleobase to afford 2’-deoxynucleosides. Enzymatic glycosylation was carried out in the presence of PyNP for thymidine (7), PNP for 2’-deoxyadenosine (32) and 2’-
Bioprocesses for the Synthesis of Nucleosides and Nucleotides
73
deoxyguanosine (1), and modified PNP for 2’-deoxycytidine (45). It is noteworthy that a 2’deoxycytidine producing enzyme was first here mentioned.
B
HO O R
HO
H Y
+
O
B
OPO32-
R
42, R=F Y=2K+
B=guanine
44, R=F B=guanine
43, R=OH Y= -
B=thymine B=adenine
7, R=OH, B=thymine (Thd) 32, R=OH, B=adenine (2'-dA)
B=guanine
1, R=OH, B=guanine (2'-dG)
B=cytosine
45, R=OH, B=cytosine (2'-dC)
Figure 20.
N-Deoxyribosyltransferases (DRTases) DRTases, because of the stereospecificity of the glycosyl transfer, may represent an alternative to nucleoside phosphorylase for enzymatic synthesis of nucleoside analogues. In spite of their strict specificity for 2’-deoxynucleosides (or 2’,3’-dideoxynucleosides), Ndeoxyribosyltransferases have a broad specificity for the acceptor bases (both pyrimidines and purines) with different substitutions. N-Deoxyribosyltransferase from L. helveticus was used to synthesize for the first time 9(2’-deoxy-β-D-ribosyl)xanthine (46) with a yield of 80% using thymidine as donor and xanthine as acceptor [68] (Figure 21). Similarly, a crude extract from L. leichmanii was used to synthesize 1-(2’-deoxy-β-D-ribofuranosyl) imidazole-4-carboxamide (48) in moderate to good yield [69] (Figure 21). Recently, [70] by using the same enzyme, 2’-deoxyguanosine (1) was synthesized in a two step reaction: first a transglycosylation reaction between thymidine (7) and 2,6diaminopurine (DAMP) and then a deamination reaction catalyzed by adenosine deaminase (ADA) (Figure 21). The same Authors reported a successful transglycosylation reaction between thymidine (7) and cytosine to give 2’-deoxycytidine (45) with a molar yield of 50% (Figure 21). This result is noteworthy because till now, no nucleoside phosphorylase has been reported to catalyze this reaction.
Marco Terreni, Daniela Ubiali, Teodora Bavaro, et al.
74
W W N
N
N
N N
N H
O
CH3
HN
+
OH 46, W=OH, Z=OH 47, W=NH2, Z=NH2
N
ADA 1, W=OH, Z=NH2 (2'-dG)
thymine
+
HO
Z
N
HO
xanthine, W=OH, Z=OH DAMP, W=NH2, Z=NH2
O
O
Z
N
O H2NOC
OH
H2NOC
7, (Thd)
TCA
N N H
NH2
O
N
N
HO O OH
N thymine
N
N
48
N H
NH2 N O
N
HO O OH 45, (2'-dC)
Figure 21.
The synthesis of 2’,3’-dideoxynucleosides was carried out with DRTase from L. helveticus using adenine, guanine, hypoxanthine, cytosine and thymine as acceptors and various 2’,3’-dideoxynucleosides as donors. An intensive interest in the preparation of these compounds arose due to their potency as inhibitors of HIV-1 infection. Among the common bases, cytosine was the best acceptor. 2’,3’-Dideoxycytidine (52) and 2’,3’-dideoxythymidine (53) were almost equivalent as 2’,3’-dideoxyribosyl donor and were superior to the purine 2’,3’-dideoxynucleosides. [71] The substrate specificity study was successfully enlarged also to some modified purine bases (6-methylpurine, 6-methylaminopurine, 6-chloropurine, 6iodopurine, 6-chloroguanine, 2-fluoroadenine, 2-chloroadenine, 2-bromoadenine, 2methyladenine, 2-aminopurine, 2,6-diaminopurine (DAMP), 2,6-dichloropurine) that were converted in the corresponding nucleosides (54-65). Only the initial rates of conversion are reported. A summary of the 2’,3’-dideoxynucleosides synthesized by DRTase from L. helveticus is reported in Figure 22.
Bioprocesses for the Synthesis of Nucleosides and Nucleotides Y
W N N HO
X
N
N N
75
Z
O
N
HO O
O
49, W=NH2, Z=H (2',3'-ddA)
58, W=Cl, Z=NH2
52, Y=NH2, X=H (2',3'-ddC)
50, W=OH Z=NH2 (2',3'-ddG)
59, W=NH2, Z=F
53 Y=OH, X=CH3 (2',3'-ddT)
51, W=OH, Z=OH (2',3'-ddI)
60, W=NH2, Z=Cl
54, W=CH3, Z=H
61, W=NH2, Z=Br
55, W=NHCH3, Z=H
62, W=NH2, Z=CH3
56, W=Cl, Z=H
63, W=NH2, Z=H
57, W=I, Z=H
64, W=NH2, Z=NH2 65, W=Cl, Z=Cl
Figure 22. 2’,3’-Dideoxynucleosides (49-65) obtained through transglycosylation catalyzed by DRT from L. helveticus.
3. Hydrolases Nucleosides synthesis by modification of the sugar moiety requires the use of protectiondeprotection reactions to ensure stereo-, chemo- and regioselectivity [2, 3]. Enzymecatalyzed deacylation has been applied as a method to selectively remove one or more acyl groups from polyacylated nucleosides. These substrates can be easily prepared by acylation of natural nucleosides or directly obtained by chemical glycosylation of the bases. Different results can be obtained depending on the substrates structure and on the enzyme. Also the reaction conditions can play an important role. The enzymatic deacetylation of 3’,5’-di-O-acetyl-2’-deoxyuridine (66a) gave the 3’deprotected product. Immobilized Candida Rugosa Lipase (CRL) and Pseudomonas Cepacia Lipase (PCL) catalyze the enzymatic hydrolysis of 66a to obtain, in 65% yield (in 4 hours) and 59% yield (in 24 hours), 5’-O-acetyl-2’-deoxyuridine (67a) (Figure 23). In the first case, also 3’-O-acetyl-2’-deoxyuridine (68a) was obtained in relevant quantity. [72] When the lipase from Porcine pancreas (PPL) was used in fully aqueous medium for deacylation of 3’,5’-di-O-acetylthymidine (66b, Figure 13), the 5’-O-acetyl group was selectively attacked leading to 3’-O-acetylthymidine (67b) in very high yield [73]. In contrast, CRL catalyzed the hydrolysis of the 3’-ester three-fold faster than the 5’-ester (Figure 23) giving a mixture of products 67b and 68b. [73] Better results have been reported by using this enzyme after immobilization. In this case, diacetylated thymidine (66b) was converted into (67b) in very high yield (90%). [74] This compound can be easily purified in multigram scale and is reported to be an useful intermediate for the synthesis of AZT [7], anti HIV agent. [8] Also the lipase from Pseudomonas fluorescens (PFL) [74, 75] was found
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to catalyze the regioselective hydrolysis at the secondary hydroxyl group. In particular, starting from different 3’,5’-di-O-acyl-2’-deoxy pyrimidine nucleosides (66 c-g), PFL affords good yields of products (67 c-g). A different selectivity was observed with the protease from B. subtilis (subtilisin) that catalyzes the hydrolysis at the primary group yielding products (68 c-g) (Figure 23) [75]. Y X
N N
O
ROCO O
Y X
N N
OH 67 a-f
O
ROCO
Y
O
X
N
OCOR 66 a-f
N
O
HO O
a b c d e f g
R =Ac, Y= OH, X = H R =Ac, Y= OH, X = Me R =C 5H 11, Y= OH, X = H R =C5 H11, Y= OH, X = Me R =C5 H11, Y= OH, X = F R =C 5H 11 , Y= OH, X = Br R =C5 H11, Y= OH, X = CF3
OCOR 68 a-f
Figure 23.
Candida antartica-Lipase B (CAL-B) catalyzes the alcoholysis of 3’,5’-di-O-acetyl-2’deoxy-nucleosides giving the corresponding 3’-O-acetyl-2’-deoxy-nucleosides in yields from 50 to 96%. [76] The alcohol employed in the biotransformation affected the rate of the enzymatic reaction and the yield of the 3’-O-acetylated products and, in all cases, a very good regioselectivity was observed. The use of alcohol both as reagent and solvent allows the solubilization of very high amounts of substrate. However, hydrolysis performed in organic solvents by using lipases usually proceeds very slowly. Subtilisin, Protease N, PPL and different microorganisms (used as whole cells) in fully aqueous medium are reported to catalyze the selective hydrolysis of the 5’-position of purine and pyrimidine tri-O-acylated esters (69 a-b) to give 2’,3’-di-O-acylribonucleosides (70 a-b). [77, 78] In particular, (Figure 14) subtilisin selectively hydrolyzes the 5’-position of 2’,3’,5’tri-O-acetyluridine (69a) giving almost quantitative yield of 2’,3’-di-O-acetyluridine (70a). This reaction is carried out in a mixture of DMF/phosphate buffer in 16 hours. It was found that PPL also catalyzes the deacetylation of 5a, but resulted in a lower selectivity and
Bioprocesses for the Synthesis of Nucleosides and Nucleotides
77
reaction rate. [74, 77, 78] Good yields of 70 a-b were also obtained by using CRL and PFL. [74] Compounds 70a can be successfully used as intermediates for the synthesis of uridine 5’-monophosphate (UMP) [74], a compound largely used as food additive [11]. Y
Y
X
X
N N
O
AcO
N
O
HO O
a b c d e
N
O
OAc OAc
OAc OAc
69 a-e
70 a-e
Y = OH, Y = NH2 , Y = OAc, Y = OH, Y = NH2 ,
X=H X=H X= H X=F X=F
Figure 24.
Good results in the selective deprotection of acetylated cytidine derivatives have been obtained by using CAL-B in alcoholysis. 2’,3’,5’-Tri-O-acetylcytidine 4b and 4-N-acetyl2’,3’,5’-tri-O-acetylcytidine (69c) were selectively converted into the corresponding nucleosides (70 b-c) (Figure 24) [76] bearing a free hydroxyl group in C-5’ position. Recently, regioselective enzymatic hydrolysis has been exploited for the preparation of 2’,3’di-O-acetyl-5-fluoro-pyrimidine (Figure 14). In particular, 2’,3’,5’-di-O-acetyl-5fluorouridine (69d) was subjected to an efficient regioselective enzymatic deprotection to give 70d catalyzed by immobilized PFL (yield 94% in 2 hours). [72, 74] Also enzymatic deacetylation of 2’,3’,5’-tri-O-acetyl-5-fluorocytidine by immobilized enzymes (CRL and Protease N) afforded a selective hydrolysis of the acetoxy group in the C-5’ position with a yield of 75-95%. [79] Compounds 70d and 70e are reported to be useful intermediates for the synthesis of Doxifluridine (74) and Capecitabine (79), respectively. Also deacylation of the peracetylated purine nucleosides 71 a-d was performed in DMF/phosphate buffer medium by using subtilisin as catalyst [77, 78] allowing to obtain the corresponding products 72 a-d selectively deacylated in C-5’ position (Figure 25). Efficient C-5’ deprotection catalyzed by CRL or PFL in CH3CN/buffer mixture has been also reported. [74] The presence of organic cosolvents in these reactions is required to ensure the complete solubilization of the substrates and to have higher reaction rates compared with the reactions performed by the free enzyme.
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B
B
AcO
HO O
O
OAc OAc
OAc OAc
71 a-d
72 a-d
a B = Adenine b B = Guanine c B = Hypoxanthine d B = N-2-Acetylguanine
Figure 25.
Also 3’,5’-di-O-acetyl-2’-deoxy -guanosine and -adenosine (73a) and (73b) (Figure 26) are efficiently deprotected in C-5’ position [76] by CAL B alcoholysis to obtain 74a and 74b, respectively. On the contrary, the acetyl esterase (AE) from the flavedo of oranges chemoand regio-selectively removes acetyl groups from these acetylated purine ribonucleosides at the C-3’ position to give nucleosides 75a and 75b, respectively. However, if the amino groups present in the bases are protected as phenylacetamides (73c and 73d), the regioselectivity completely changes and nucleosides 74c and 74d are obtained [73]. W
W N N AcO
N
N N
Z
O OAc 73 a-d
N HO O OAc 74 a-d
W N
N N
Z
N AcO
N N
Z
O OH 75 a-b
a W = NH2 , Z = H b W = H, Z = NH2 c W = NHCOCH 2ph, Z = H d W = OH, Z = NHCOCH 2ph
Figure 26.
The hydrolysis of nucleosides protected with alkoxycarbonyl groups has been reported by using Pig Liver Esterase (PLE) and CAL-B. Complete cleavage of carbonate and carbamate functions is reported by using PLE at pH 7 and 60 °C and CAL B under milder conditions. [80] Regioselective hydrolysis is only reported to obtain 2’,3’-di-O-ethoxycarbonyluridine (77a) and 2’,3’-di-O-ethoxy-carbonylinosine (77b) by regioselective alcoholysis (Figure 27). Specifically, CAL B catalyzes the regioselective ethanolysis of the trialkoxycarbonylated nucleosides 76 a-b with benzyl alcohol in 1,4-dioxane at 30 °C [81].
Bioprocesses for the Synthesis of Nucleosides and Nucleotides B
79
B
RO
HO O
O
OR OR
OR OR
76 a-b
77 a-b
a R = COOEt, B = Uridine b R = COOEt, B = Hypoxanthine
Figure 27.
Only few results have been reported concerning the selective deprotection of acetylated arabino nucleosides. Recently, [72] it has been reported the regioselective hydrolysis of peracetylated arabinosyluridine (78a) (Figure 28) by using immobilized lipases: CRL afforded 2’,3’-di-O-acetylarabinosiluridine (79a) in 89% yield; PCL displayed a moderate activity (yield 66% after 24 hours). The hydrolysis of tri-O-acetylated esters 78b with Pig Liver Esterase (PLE) in phosphate buffer with ethanol as cosolvent leads to 2’-Omonoacetylated nucleosides 79b in high yield (Figure 28). The markedly retarded rates of hydrolysis of the 2’-O-acyl esters of these arabinonucleosides suggest that they might function as slow release lipophilic prodrugs with long serum lifetimes [73]. O X
O X
NH N
O
RO
NH N
O
RO O OAc
O OAc
OR1
OR 1
78 a,b
79 a,b
a X = H, R = R1 = OAc b X = CHCHBr, R = R 1 = OAc
a X = H, R = OH, R1 = OAc b X = CHCHBr, R = R 1 = OH
Figure 28.
It is well known that hydrolases catalyze the enantioselective hydrolysis/synthesis of a wide range of soluble or insoluble chiral esters. In this context, Wengel and co-workers [82] reported the first attempt to use biotransformations to solve the basic problem of anomer separation in nucleoside chemistry when a convergent strategy has been used. Thus, lipase catalyzed deacetylations of anomeric mixtures of peracetylated 2'deoxyribofuranosyl- thymine nucleosides (80) are shown in Figure 29. Generally, the diastereoselectivity was more pronounced in pure phosphate buffer than in phosphate buffer containing 10% DMF. Wheat Germ Lipase (WGL) and Porcine Liver Esterase (PLE)
Marco Terreni, Daniela Ubiali, Teodora Bavaro, et al.
80
catalyzed the diastereoselective deacetylation of 80 affording the pure ß-anomer thymidine (81ß) as the only completely deprotected nucleoside product [73]. O
O NH
N AcO O OAc 80 α, β
NH
O
N
O
HO O OH 81 β WGL PLE
29% yield 31% yield
Figure 29.
According to the results reported in this review, the screening of different hydrolases allows to find catalysts able to regioselectively deprotect different natural and unnatural acetylated nucleosides in crucial positions. By this enzymatic approach, a library of monohydroxy acetylated nucleosides can be prepared in good overall yields, by selecting, for each substrate, a specific enzyme suitable to ensure the most adequate activity and regioselectivity. However, the screening of different enzymes, many of them available on the market, is only the first step for the development of an efficient industrial preparative process. In fact, for the development of the process, as previous described for nucleoside phosphorylases, the engineering development of the catalyst is fundamental to achieve high performances in terms of activity and stability in the reaction conditions necessary to obtain high yields and high product concentration. Most of hydrolyses reported in literature and catalyzed by lipases have been performed by using high concentrations of fully soluble enzymes or “straight from bottle” enzyme preparations. Because of the trend of lipases to be activated (adsorbed) on hydrophobic structures [83], many of these biotransformations catalyzed by fully or partially soluble lipases in macro-aqueous media could be influenced by lipase-lipase or lipase-protein interactions. Furthermore, in this medium, the equilibrium between the close and the open active form of these enzymes is almost completely shifted towards the inactive closed form. Thus, reaction proceeds very slow and side reactions catalyzed by other enzymes present in commercially available crude extract of lipases (including proteolysis of the lipase) cannot be excluded, and could be responsible of both a low purity of the product and enzyme stability. In some cases, over-saturated substrate solutions (“oil in waters” suspensions) are used and hence soluble lipases may undergo interfacial activation on drops of substrate [84]. However, this mechanism of activation is not really applicable for water soluble substrates such as nucleosides, even if used in their acetylated form. The use of immobilized enzymes, instead of fully or partially soluble preparations, may present many advantages, mainly bearing in mind a possible scaling up of the process (easy performance and design of the bio-reactor, recovery of pure products, reuse of the biocatalyst,
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81
operational stabilization of the enzyme via immobilization techniques, etc). In order to achieve biocatalysts with a high volumetric activity, a large amount of lipase must be immobilized in the support and this makes necessary the use of porous supports. Fully dispersed immobilized lipase molecules cannot undergo lipase-lipase or lipase-protein interactions and thus also inactivation by proteolysis is avoided. In this case, even using oversaturated substrate suspensions, the immobilized lipase molecules (because placed inside a porous support) cannot undergo interfacial activation [85] and can only act on the fraction of soluble substrate in the reaction medium and, therefore, able to penetrate inside the porous structure of the support. It is also important to remark that different immobilization protocols (involving different areas of the protein or promoting different degrees of enzyme rigidity) could induce a different shift in the equilibrium between the closed and the open structure of the lipase, as well as a different shape of the active site of the open structure of immobilized lipases. This may yield catalysts with different activity, but also different kinetic properties such as regioand enantio- selectivity [86]. In particular, covalent immobilization of lipases affords inactive catalysts being the enzyme immobilized in the closed form and/or because of the flexibility reduction of the enzyme structure necessary to shift the equilibrium from the closed to the open structure. On the contrary, adsorption of lipases on hydrophobic support ensures the simultaneous immobilization, activation and purification of these enzymes. In fact, the open active form is stabilized by the adsorption of the enzyme on the support surface by interaction of the hydrophobic areas near the active site. Furthermore, if the adsorption on hydrophobic supports is performed at a very low ionic strength, the process becomes very selective for the lipase immobilization, allowing the simultaneous purification of these enzymes from the crude extracts [85, 87]. For example, the use of lipases as catalysts in the regioselective hydrolysis of peracetylated sugars, including ribose, has been reported. [88] In this case, immobilization of these enzymes by hydrophobic adsorption improved activity and yields compared with covalent immobilization. Also in the hydrolysis of acylated nucleosides, the use of immobilized lipases gives better results than those achieved by the free enzymes. For example, immobilized CRL [74] allows to obtain higher regioselectivity and yields compared with the use of free CRL [73] as reported in the hydrolysis of diacetylated thymidine (66b), and to obtain compound 68b bearing a free hydroxyl group in C-3’ position (see Figure 23). Reaction conditions: 20 mM solution of substrate in 25 mM potassium phosphate buffer and acetonitrile (10%) at pH=7; 50 units of enzyme in 35 mL. During the reaction the pH was kept constant by automatic titration. Samples of the reaction mixture were analyzed at different times by TLC and HPLC. To confirm the that improved results, compared with the free enzyme, can be obtained using lipases immobilized on hydrophobic supports, we have performed (unpublished results) the hydrolysis of triacetylated uridine (69a) to obtain the compound 70a bearing a free hydroxyl group in C-5’, by comparing the free CRL with the catalyst obtained after immobilization of this enzyme through adsorption on hydrophobic support (Figure 30). Also in this case, the immobilization improves the performances of the lipase, compared with the
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free enzyme. In fact, immobilized CRL gives about 90% yield (in 24 hours) while by free CRL the monodeprotected compound was obtained in 30% yield.
Figure 30. Hydrolysis of 69a catalyzed by the free and immobilized lipase from Candida rugosa.
Nucleosides selectively deprotected in different positions can be used for developing new synthetic routes for the preparation of various modified nucleosides. For example, modification of the sugar moiety can be performed directly starting from uridine. This natural and cheap starting material can be selectively modified in C-5’, after chemical fully acetylation, as a consequence of the very high selectivity of immobilized PFL or CRL. In fact, 2’,3’,5’-tri-O-acetyluridine (69a) was subjected to an efficient and complete regioselective enzymatic hydrolysis to give 70a. This protected nucleoside bearing only one free hydroxyl group can be directly used without purification as intermediate for functionalization. O
O NH
N HO O
NH
O
--
(C2 H5 )3 PO4 /POCl3
N
O
O3 PO O
70% OAc OAc 70a
OH OH 82, UMP (5'-monophosphate uridine)
Figure 31.
It has been reported the use of this intermediate for the preparation of the corresponding 5’-monophosphate used both in pharmaceutical and food industry. [74] The phosphorylation of 70a (Figure 31) was performed by phosphorus oxycloride (POCl3) in trialkyl phosphates (TAP) to directly give the deprotected nucleoside 5’-monosphosphates after the work-up. In
Bioprocesses for the Synthesis of Nucleosides and Nucleotides
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fact, uridine 5’-monophosphate (82, UMP) was obtained by a chemo-enzymatic two-steps process starting from per acetylated uridine (69a). Starting from the intermediate 70a, also a new method for the synthesis of Doxifluridine (87) has been proposed. [72] In fact, 70a was deoxygenated and deprotected affording 5’deoxyuridine (86) (Figure 32). This compound was used for the enzymatic transglycosylation (pyrimidine/pyrimidine) with 5-fluorouracil, catalyzed by immobilized UP from B. subtilis [53, 54], to yield Doxifluridine (87) in about 40% of conversion (not optimized).
Figure 32.
This process presents several advantages compared with the classical chemical process used for the synthesis of this compound [13]. This new synthetic route for Doxifluridine drastically reduces the number of hazardous chemical steps because glycosylation of the fluorinated uridine is performed only in the last step and by an enzymatic reaction. Furthermore, the requirement of the protection and deprotection of reactive groups is sensitively reduced and two synthetic steps are performed in fully or almost fully aqueous medium. Alternatively, fully acetylated 5-fluorouridine or 5-fluorocytidine, prepared by chemical glycosylation with peracetylated ribose can be used, after selective enzymatic hydrolysis in C-5’ catalyzed by immobilized CRL, as intermediates for the synthesis of Doxifluridine [74] and Capecitabine [79].
4. Enzymatic Phosphorylation Nucleotides are phosphoric acid esters of nucleosides in which the phosphoric acid is esterified to one of the free pentose hydroxyl groups. Nucleotides occur in free form in
84
Marco Terreni, Daniela Ubiali, Teodora Bavaro, et al.
significant amounts in a variety of cell types. They are also formed on partial hydrolysis of nucleic acids, particularly by the action of a class of enzymes called nucleases. The major function of the 2'-deoxyribose nucleotides is in DNA. The ribonucleotides are the monomeric units of RNA but also serve in most other cellular and metabolic functions of nucleotides. The phosphoryl group of nucleotides is most commonly esterified to the C-5' hydroxyl of the pentose. The number of phosphate groups attached is indicated by a mono-, di- or tridesignation. Nucleotides (NMPs) are often used as food additives and as pharmaceutical intermediates. Among them, 5’-IMP and 5’-GMP are important because they have a characteristic taste and are used as flavor enhancers in various foods. Human milk was investigated by Sugawara et al. [10] and three nucleosides (cytidine, uridine and adenosine), and six nucleotides: cytidine 5′-monophosphate (5′-CMP), uridine 5′-monophosphate (5′UMP), adenosine 5′-monophosphate (5′-AMP), guanosine 5′-monophosphate (5′-GMP), inosine 5′-monophosphate (5′-IMP) and cytidine 5′-diphosphate (5′-CDP) were quantified. The Authors suggested that nucleosides and nucleotides found in human milk might play some important roles in the development of infants. On the other hand, the total nucleotide content has been found to be much lower in bovine milk and bovine milk-based infant formula than in human milk, although orotic acid is present in significant quantities in cow's milk and is a precursor of pyrimidines. [11] In order to ensure that infants gain the same nutritional benefits from infant milks and formulae as those from human milk, nucleotide supplementation is permitted by regulatory agencies in the USA, Europe and Japan, as well as some other countries. These countries have accepted the safety of the practice of nucleotide addition to infant formulae, because these compounds are present in human milk and, thus, there is a high likelihood that they are of benefit to infants. Since there are two or more free hydroxyl groups in nucleosides, the phosphate group of nucleotides can potentially occur in more than one position on the sugar ring. In the case of deoxyribonucleotides, there are only two possible positions that can be esterified with phosphoric acid, namely, the 3' and 5' positions. In the case of ribonucleotides, the phosphate group may be at the 2', 3', or 5' position. As a consequence, the chemical synthesis is usually performed by using a protecting groups strategy. NMPs can be also obtained by microbial synthesis or by isolation from hydrolysates of nucleic acids, which are the main sources of these compounds [89]. Purine nucleosides such as inosine [90] and guanosine [91] can be produced efficiently by fermentation, and phosphorylation of nucleosides is a very efficient process for the large-scale production of 5’nucleotides.
4.1. Kinases Deoxyribonucleoside kinases catalyze the phosphorylation of deoxyribonucleosides (dN) to the corresponding deoxyribonucleoside monophosphates (dNMP). They are the key enzymes in the salvage of deoxyribonucleosides originating extracellularly from food/medium or from intracellular breakdown of DNA [92].
Bioprocesses for the Synthesis of Nucleosides and Nucleotides RO O RO
RO
OR1
Base
Glycosylation
Deprotection
O
OR
RO
85
HO
Base O
OR
HO
OH
R=Bz, R1 =Ac
Protection
HO
Base
Functionalization
OR
Deprotection
O RO
X HO
Base O Y
Figure 33.
Similarly, nucleoside monophosphate kinases (ATP/nucleoside monophosphate phosphotransferases) are well known enzymes able to catalyze the phosphorylation of nucleoside monophosphates to the corresponding diphosphates using the terminal phosphate group of ATP Finally, nucleoside diphosphate kinases (ATP/nucleoside diphosphate phosphotransferases) catalyze instead the phosphorylation of nucleoside diphosphates also using the terminal phosphate group of ATP (Figure 33). Deoxyribonucleoside kinases are the most studied catalysts for nucleotides synthesis. These enzymes can be in fact useful for preparation of NMPs with high regioselectivity, avoiding any protection of the base and sugar moiety and in fully aqueous medium due to the very high water solubility of the substrates. However, the very high selectivity of these enzymes strongly limits the applications at a preparative scale. Particularly, mammal cells have four deoxyribonucleoside kinases with overlapping specificities: cytoplasmic thymidine kinase 1 (TK1) which phosphorylates only thymidine, mitochondrial thymidine kinase 2 (TK2) which phosphorylates deoxycytidine and thymidine, cytoplasmic deoxycytidine kinase (dCK) which phosphorylates deoxyadenosine, deoxyguanosine and deoxycytidine, and mitochondrial deoxyguanosine kinase (dGK) which phosphorylates only purine deoxynucleotides, deoxyadenosine and deoxyguanosine. All four human kinases have been cloned and expressed in bacteria [93]. Although one of the cellular kinases, deoxycytidine kinase (dCK), has been shown to phosphorylate several L-nucleosides (used as efficient antiviral agents) [94], generally, the high specificity of this enzyme makes necessary the study of a suitable catalyst for each nucleoside used as substrate. Furthermore, kinases require the use of expensive nucleoside triphosphates as phosphate donor (for example ATP that is converted into ADP). This problem can be solved by using recycling enzymes (other kinases) that, starting from cheap reagents used as phosphate donors, catalyze the recycle of ADP, allowing the use of ATP only in catalytic amounts. However, the simultaneous use of different enzymes makes difficult the development of efficient processes, involving immobilized enzymes, for preparative purpose. In fact, no example of immobilization of deoxyribonucleoside kinases have been reported, whereas only few examples of application of these enzymes at a preparative scale are available in literature.
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The problem of the extreme selectivity of deoxyribonucleoside kinases has been partially solved by the research of natural sources for enzymes with a wider range of selectivity. For example, in the fruit fly, only a single deoxyribonucleoside kinase exists (DmdNK, EC 2.7.1.145), capable of phosphorylating all four natural substrates. The multisubstrate deoxyribonucleoside kinase of Drosophila melanogaster (Dm-dNK) was found to be the fastest known deoxyribonucleoside kinase enzyme. [95] Its sequence is related to three human kinases and to herpes simplex virus type-1 thymidine kinase. Dm-dNK phosphorylates both purine and pyrimidine deoxyribonucleosides and nucleoside analogues although it has a preference for pyrimidine nucleosides. Recently, a recombinant kinase from D. melanogaster has been reported in a patent application [96] as catalyst for the synthesis of d-GMP 88 and d-AMP 89 in around 80% yield (Figure 34). R1
R1 N N HO O
N
N N
N
R2
HO O HO P O
R
OH
ATP 1 32 30
ADP
R=H R1=OH R2=NH2 R=H R1=NH2 R2= H R=OH R1=NH2 R2= F
O
N N
R2
R
OH
88 89 90
Figure 34.
The DNA sequence encoding the kinase as well as a procedure for preparation of the enzyme and its use during the synthesis of nucleoside monophosphates have been described. According to the patent, this enzyme could be applied also in the synthesis of the different nucleoside monophosphates, deoxynucleoside monophosphates, dideoxynucleoside monophosphates as well as other sugar- and base-modified nucleoside monophosphates. Recently, [97] the cloning of three genes from Dictyostelium discoideum and the characterization of the corresponding thymidine kinase (DdTK1), deoxyadenosine kinase (DddAK), and deoxyguanosine kinase (DddGK) have been described. These enzymes have very narrow substrate specificities. DdTK1 phosphorylates only thymidine. DddAK is highly specific for deoxyadenosine but can also poorly phosphorylate thymidine. Fludarabine (F-araA) is an antimetabolite pro-drug used in the treatment of B-cell chronic lymphocytic leukemia. [12] As an adenosine analog, 2-fluoroadenosine 30 is poorly water-soluble, therefore, to increase the solubility it is used in the monophosphorylated form (F-araA). This compound has also been considered for a synthesis mediated by kinases. Particularly, despite the high specificity reported for DddAK [97] this enzyme resulted also very active for phosphorylation of 30, to obtain F-araA (90) by using ATP or GTP as phosphate donors (Figure 23). Thus, this enzyme could be a potential catalyst for production
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of Fludarabine that today relies on traditional organochemical synthesis leading to high production cost and use of polluting solvents [98]. The main problem related to the use of kinases in the synthesis of mucleotides is the efficient purification of the target product from other mono-, di- and tri- phosphate esters. A number of chromatographic procedures have been described which ensure a reliable separation of nucleotide mixture components at the end of the reaction. In particular, such are chromatographic fractionations on DEAE cellulose, Dowex 1×2, Aminex A, PAP. At acidic pH values, nucleotides are fractionated in a formate system (suitable for liophilization) and at acidic and alkaline pH values fractionation is performed in a chloride system. At pH > 8.0, ammonium bicarbonate can also be used [99]. To fractionate nucleotide mixtures at a preparative scale, an anion exchanger Dowex 1×2 or its analog AB 17×2 are the best. These anion exchangers have high capacity and allow to perform chromatography at a sufficiently high rate of elution. Fractionation of nucleotide mixtures using the above mentioned anion exchangers results in the most complete separation of nucleoside monophosphates from nucleoside di- and tri-phosphates and also the best separation of nucleotides themselves. A new, simple, and ingenious method for enzymatic synthesis of deoxy- and ribonucleoside-5′-triphosphates (dNTP and NTP, respectively) has been developed by Bochkov. [100] The method includes the following stages: hydrolysis of DNA with DNase and immobilized S1-nuclease, phosphorylation of the resulting deoxy- and ribonucleoside-5′monophosphates (dNMP and NMP) with nucleotidyl kinase from E. coli, and purification by chromatography of the synthesized dNTP and NTP. dNMP was then further phosphorylated by using an ATP-regenerating system based on acetokinase from E. coli and lithium acetylphosphate.
4.2. Nucleoside Phosphotransferases Microbial and plant nucleoside phosphotransferases (EC 2.7.1.77; NPase) are also widely applied as biocatalysts for the 5’-monophosphorylation of nucleosides. As distinct from nucleoside kinases, these enzymes have a broader specificity with respect to both the phosphate donors and acceptors, and they can accept the low-energy monoesters of phosphoric acid such as, for example, natural NMP and p-nitrophenylphosphate (p-PNP) as substrates. A method to transform nucleosides into 5’-monophosphates using nucleoside phosphotransferase from Erwinia herbicola as a wet paste of bacterial cells has been reported. [101] This method is based on the shift in the equilibrium state of the reaction to the formation of desired product due to its precipitation by Zn2+. Interestingly, by using the wet paste of bacterial cells, the synthesis of Fludarabine (90), and many other arabinonucleoside-5’-monophosphates (92-96, Figure 35) have been reported with yields between 41% and 59% (mol). Similarly, under optimal conditions, adenosine (6) is converted into AMP (97) in 83% yield (Figure 34).
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Marco Terreni, Daniela Ubiali, Teodora Bavaro, et al. B HO O
R
OH R' 28 R=OH R'=H 29 R=OH R'=H 91 R=OH R'=H 23 R=OH R'=H 34 R=OH R'=H 6 R=H R'=OH 33 R=H R'=OH
B
HO O HO P O
O
R
OH R' B= Adenine B= 2-Amino adenine B= Guanine B= Uracil B= Cytosine B= Adenine B= Hypoxanthine
92 93 94 95 96 97 98
Figure 35.
4.3 Acid Phosphatases/Phosphotransferases and Nonspecific Phosphohydrolases A mutated acid phosphatase/phosphotransferase (AC/ATase EC 3.1.3.2) from Morganella morganii catalyzes the nucleoside phosphorylation reaction using pyrophosphate (PPi) as a donor. Yasuhiro Mihara et al [102] patented a method for producing nucleoside-5'phosphate esters inexpensively and in high yields by phosphorylating a nucleoside with a phosphate group donor by using acid phosphatases having an increased affinity for the nucleoside and/or an increased temperature stability at a pH of 3.0 to 5.5. This method can be used in the synthesis of 5’-IMP (98) from inosine (33) by E. coli overproducing AP/PTase. The productivity in 5’-IMP is around 45g/l in 2.5 hours (Figure 35). Bacterial nonspecific acid phosphohydrolases (NSAPs) are secreted enzymes, produced as soluble periplasmic proteins or as membrane-bound lipoproteins, that are usually able to dephosphorylate a broad array of structurally unrelated substrates and exhibit optimal catalytic activity at acidic to neutral pH values. The term ‘NSAP’ was originally adopted to indicate bacterial enzymes which, unlike alkaline phosphatase, show optimal catalytic activity at acidic to neutral pH values and, unlike specific phosphohydrolases (e.g. 3’nucleotidases, 5’-nucleotidases, hexose-phosphatases and phytases), do not exhibit a marked substrate specificity, retaining activity towards several different and structurally unrelated phosphoesters. Bacterial NSAPs are monomeric with a Mw of 25–30 kDa. On the basis of amino acid sequence relatedness, three different molecular families of NSAPs can be distinguished, indicated as molecular class A, B and C, respectively. Members of each class share some common biophysical and functional features, but may also exhibit functional differences. [103] The first bacterial NSAPs purified and characterized in detail were the periplasmic PhoN (or nonspecific acid phosphatase I) and AphA (or nonspecific acid phosphatase II) enzymes produced by S. enterica ser. Typhimurium [104]. Nevertheless, among the considerable number of strains studied, only Providencia stuartii and Morganella morganii are able to produce a high level of acid phosphatase
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activity unlike most other enterobacterial species, which produce only low to moderate levels of acid phosphatase activity. The potential utility of some class A bacterial NSAPs in the sector of applied microbiology and biotechnology has not been fully exploited.
5. Conclusion Bioconversions here described have the potential to shorten the preparation processes by reducing or eliminating the need of protecting groups, avoiding the formation of regio- and stereo-isomers and decreasing production costs. Moreover, since the reactions are carried out mostly in water solutions, environmental pollution is sensitively reduced. The most studied reactions are based on sugar exchange between a nucleoside donor and a purine or pyrimidine base acceptor catalyzed by NPs, or on regio-selective deprotection of sugar moiety, mediated by hydrolases. Also the use of regio-selective enzymes for phosphorylation is becoming an attractive tool for nucleoside synthesis. The selection of the biological source plays a crucial role in determining the substrate specificity. The numerous examples here reported, however, clearly showed that a large “enzyme toolset” is available for the synthesis in high yield of natural and unnatural nucleosides. Besides, the enzyme immobilization techniques can further assist the successful development of such bioconversions at a preparative-large scale as they can enhance the catalyst stability, its recovery and re-use with obvious advantages. In particular, if the immobilization occurs through the formation of covalent linkages between the enzyme and the solid support it can ensure that no protein contaminants are released in the reaction mixture, preserving the quality of the final compound. In the case of substances intended for human use (e.g. drugs or food ingredients) this is a prerequisite for regulatory approval and, therefore, cannot be underestimated. In this context many effort should be made to propose a set of catalyst suitable for industrial application. For that concerning phosphorilases, in fact, only few example of efficient immobilization are available in the literature, and the development of stable catalyst from others NPs with different selectivity should be an important future development. Similarly, for that concerning hydrolases, efficient catalyst have been developed, for regio-selective hydrolysis of per-acylated nucleosides, by immobilization of lipases by hydrophobic adsorption on solid support. However these lipase preparations could be not suitable to ensure a complete stability of the catalyst in drastic condition and, thus to avoid also the release of the protein in the reaction medium. Consequently, for these reactions, an interesting topic, could be the study and the use of immobilized esterases, being these enzymes efficiently immobilized by covalent linkage. Finally, but not last, the study of efficient immobilization procedure also for phosphorilating enzymes should be also adequately considered in the future.
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References [1] [2] [3]
[4]
[5] [6] [7] [8]
[9]
[10] [11]
[12] [13] [14]
[15] [16] [17]
[18] [19]
Komatsu, H., Awano, H., Tanikawa, H., Itou, K., and Ikeda, I. Nucleosides, Nucleotides & Nucleic Acids, 2001; 20: 1291. Vorbruggen, H. and Ruh-Pohlenz, C. “Synthesis of nucleosides” in Organic Reactions (L.A. Paquette Ed.), Wiley, New York, 2000, Vol. 55. Niedballa, U. and Vorbruggen, H. J Org Chem. 1974; 39: 3654; b) Jungmann, O. and Pfleiderer W. Tetrahedron Lett. 1984; 25: 5017; c) Kawakami, H., Matsushita, H., Naoi, Y., Itoh, K., and Yoshikoshi, H. Chem Lett. 1989; 235. ICH Topic Q 3 A (R2) Note for guidance on impurities testing: impurities in new drug substances, EMEA, October 2006; b) ICH Topic Q 3 B (R2) Note for guidance on impurities in new drug product, EMEA, June 2006. General monograph of European Pharmacopoeia 6.0 2008 “Substances for pharmaceutical use” Vol 1: 703. ICH Topic Q 3 C (R3) Note for guidance on impurities: residual solvents, EMEA, March 1998. Chen, B.C. and Quinlan,S.L. 1995 US 5466787. Barre-Sinoussi, F., Chermann, J.C., Rey, F., Nugeyre, M.T., Chamaret, S., Gruest, J., Dauguet, C., Axler-Blin, C., Vezinet-Brun, F., Rouzioux, C., Rozenbaum, W., and Montagnier, L. Science, 1983; 220: 868. Montgomery, J.A. and Shortnacy, A.T. 1982 US Patent 4,357,324; b) Ikemoto, T., Haze, A., Datano, H., Kitamoto, Y., Ischida, M., and Nara, K. Chem Pharm Bull. 1995; 43: 210. Sugawara, M., Sato, N., Nakano, T., Idota, T., and Nakajima, I.J Journal of Nutritional Science and Vitaminology, 1995; 414: 409. Thorell, L., Sjoeberg, L.B. and Hernell, O. Pediatric Research, 40: 845; b) Damkjaer, D.L., Petersen, M., and Wengel, J. Nucleosides Nucleotides, 1994; 13: 1801; c) Jyonouchi, H.J. Nutr. 1994; 124: 138S; d) Van Buren, C.T.; Kulkarni, A.D., and Rudolph, F.B.J. Nut, 1994; 124: 160S. Keating, M.J., Kantarjian, H., Talpaz, M., Redman, J., Koller, C., Barlogie, B., Velasquez, W., Plunkett, W., Freireich, E.J., and McCredie, K.B. Blood, 1989; 74: 19. Cook, A.F. 1978 US Patent 4,071,680. Fujiu, M., Ishitsuka, H., Masanori, M., Umeda, I., and Yokose, K. 1990 US Patent 4,966,891; b) Arasaki, M., Ishitsuka, H., Kuruma, I., Masanori, M., Murasaki, C., Shimma, N., and Umeda, I. 1995 US Patent 5,472,949. Longley, D.B., Harkin, D.P. and Johnston, P.G. Nature Reviews Cancer, 2003; 3: 330. Di Costanzo, F., Sdrobolini, A. and Gasperoni, S. Critical Reviews in Oncology/Hematology, 2000; 35: 101. Wong, C.H. and Whitesides, G.M. “Enzymes in synthetic organic chemistry,” Pergamon, Oxford, 1994; b) Prasad, A.K., Trikha, S., Parmar and V.S. Bioorg Chem. 1999; 27: 135. Tao, J., Zhao, L. and Ran, N. Org. Proc. Res. & Dev. 2007; 11: 259. General monograph of European Pharmacopoeia 6.0 2008, “Product of fermentation” Vol 1, 693.
Bioprocesses for the Synthesis of Nucleosides and Nucleotides
91
[20] Drauz, K. and Waldmann, H. “Enzyme catalysis in organic synthesis: a comprehensive handbook”, VCH, Weinheim, 2002; b) Faber, K. “Biotransformations in organic chemistry”, 4th Ed. Springer, Berlin, 2000. [21] Klibanov, A.M. Adv Appl Microbiol. 1983; 29: 1; b) Gupta, M.N. Biotechnol Appl Biochem. 1991; 14: 1; c) Gianfreda, L. and Scarfi, M.R. Mol Cell Biochem. 1991; 100: 97; d) Fernández-Lafuente, R., Rodríguez, V., Mateo, C., Penzol, G., Hernández-Justiz, O., Irazoqui, G., Villarino, A., Ovsejevi, K., Batista, F., and Guisán, J.M. J. Mol. Cat. B 1999; 7: 181; e) Fernández-Lafuente, R.H., Jústiz, O., Mateo, C., Terreni, M., Fernández-Lorente, G.,. Moreno, M.A., Alonso, J., García-López, J.L., and Guisan, J.M. Biomacromolecules, 2001; 2: 95. [22] Pugmire, M.J. and Ealick, S.E. Biochem J. 2002; 361: 1. [23] Krenitsky, T.A., Mellors, J.M. and Barclay, R.K. J Biol Chem. 1965; 240: 1281. [24] Pugmire, M.J. and Ealick, S.E. Structure, 1998; 6: 1467. [25] Walter, M.R., Cook, W.J., Cole, L.B., Short, S.A., Koszalka, G.W., Krenitsky, T.A., and Ealick, S.E. J Biol Chem. 1990; 265: 14016. [26] Zimmerman, M. and Seidenberg, J. J Biol Chem. 1964; 239: 2618; b) Zimmerman, M. and Seidenberg, J. J Biol Chem. 1964; 239: 2622. [27] Gallo, R.C. and Breitman, T.R. J Biol Chem. 1968; 243: 4936. [28] Schwartz, M. Eur J Biochem. 1971; 21: 191. [29] Avraham, Y., Grossowicz, N. and Yashphe, J. Biochim Biophys Acta, 1990; 1040: 287. [30] Krenitsky, T.A. and Tuttle, J.V. Biochim Biophys Acta, 1982; 703: 247. [31] Degranges, C., Razaka, M. and Bricaud, H. Biochim Biophys Acta, 1981; 654: 211. [32] Cardinaud, R. Methods Enzymol. 1978; 5: 446. [33] Kilstrup, M., Hammer K., Jensen, P.R., and Martinussen, J. FEMS Microbiol Rev. 2005; 29: 555. [34] Miyamoto, Y., Masaki, T. and Chohnan, S. Biochim Biophys Acta, 2007; 1774: 1323. [35] Kaminski, P.A. J Biol Chem. 2002; 277: 14400. [36] Steenkamp, D.J. Eur J Biochem. 1991; 197: 431; b) Steenkamp, D.J. and Halbich, T.J. Biochem J. 1992; 287: 125. [37] Holguin, J. and Cardinaud, R. Eur J Biochem. 1975; 54: 505. [38] Armstrong, S.R., Cook, W.J., Short, S.A., and Ealick, S.E. Structure, 1996; 4: 97. [39] Anand, R., Kaminski, P.A. and Ealick, S.E. Biochemistry, 2004; 43: 2384. [40] Bzowskam A., Kulikowska, E. and Shugar, D. Pharmacol Therap. 2000; 88: 349. [41] Krenitsky, T.A., Koszalka, G.W. and Tuttle, J.V. Biochemistry, 1981; 20: 3615. [42] Krenitsky, T.A. Mol Pharm. 1967; 3: 526; b) Krenitsky, T.A. J Biol Chem. 1968; 243: 2871. [43] Chae, W.G. Biotech and Bioprocess Engin. 1999; 4: 17. [44] 44) Betbeder D, Heath CM, Hutchinson DW. Nucleosides Nucleotides 1991;10: 465. [45] Hutchinson, D.W. Trends Biotechnol. 1990; 8: 348; b) Hanrahan, J.R., Hutchinson and D.W. J Biotechnol 1992; 23: 193; c) Utagawa, T. J Mol Cat B: Enzym. 1999; 6: 215. [46] Bestetti, G., Cali', S., Ghisotti, D., Orsini, G., Tonon, G., and Zuffi, G. 2000 WO 00/39307 A2; b) Yokoseki, K., Tsuji, H. and Izawa, K. 1999 EP 916735 A1; c) Kojima, E., Yoshioka, H., Fukinbara, H., and Murakami, K. 1990 US 5563049 A; d) Yokozeki,
92
[47] [48] [49] [50] [51] [52]
[53]
[54] [55] [56] [57] [58] [59] [60] [61] [62] [63] [64]
[65] [66] [67] [68] [69]
Marco Terreni, Daniela Ubiali, Teodora Bavaro, et al. K., Shirae, H., Shiragami, H., Irie, Y., Yasuda, N., Otani, M., and Tanabe, T. 1989 US 4835104 A. Trelles, J.A.., Valino, A.L., Runza, V., Lewkowicz, E.S., and Iribarren, A.M. Biotechnol Lett. 2005; 27: 759. Barai, V.N., Zinchenko, A.I., Eroshevskaya, L.A., Kalinichenko, E.N., Kulak, T.I., and Mikhailopulo, I.A. Helv Chim Acta 2002; 85:1901. Hennen, W.J. and Wong, C.H. J Org Chem. 1989; 54: 4692. Hatano, A., Harano, A. and Kirihara, M.Chem Letters, 2006; 35: 232. Mahmoudian, M. Biocat and Biotransf. 2000; 18: 105. Zuffi, G., Ghisotti, D., Oliva, I., Capra, E., Frascotti, G., Tonon, G., and Orsini, G. Biocat and Biotransf. 2004; 22: 25; b) Tonon, G., Capra, E., Orsini, G., and Zuffi, G. 2004 EP 1439220 A1. Rocchietti, S., Ubiali, D., Terreni, M., Albertini, A.M., Fernandez-Lafuente, R., Guisan, J.M., and Pregnolato, M. Biomacromolecules, 2004; 5: 2195; b) Pregnolato, M., Terreni, M., Albertini, A.M., Guisan, J.M., Fernandez Lafuente, R., and Frigerio, M, 2003 WO 03/008619 A1. Ubiali, D., Rocchietti, S., Scaramozzino, F., Terreni, M., Albertini, A.M., FernandezLafuente, R., Guisan, J.M., and Pregnolato, M. Adv Synth Catal. 2004; 346: 1361. Hori, N., Watanabe, M., Sunagawa, K., Uehara, K., and Mikami, Y. J Biotechnol. 1991; 17: 121. Goldstein, L., Levin, Y. and Katchalski, E. Biochemistry, 1964; 3: 1913. Mateo, C., Abian, O., Fernandez-Lafuente, R., and Guisan, J.M. Biotechnol Bioeng. 2000; 68: 98. Torres, R., Mateo, C., Fuentes, M.; Palomo, J.M., Ortiz, C., Fernandez-Lafuente, R., Guisan, J.M., Tam, A., and Daminati, M. Biotechnol Prog. 2002; 18: 1221. Mikami, Y., Matsumoto, S., Yoshinaka, S., Sunaga, Y., and Hasegawa, A. 2000 US 00/6017736 A. Zinchenko, A.I., Barai, V.N., Bokut, S.B., Kvasyuk, E.I., and Mikhailopulo, I.A. Appl Microbiol Biotechnol. 1990; 32: 658. Barai, V.N., Zinchenko, A.I., Eroshevskaya, L.A., Zhernosek, E.V., De Clercq, E., and Mikhailopulo, I.A. Helv Chim Acta, 2002; 85:1893. Ciuffreda, P., Loseto, A. and Santaniello, E. Tetrahedron. 2002; 58: 5767. Gupta, M. and Nair, V. Collect Czech Chem Commun. 2006; 71: 769. Exall, A.M., Jones, M.F., Mo, C.L., Myers, P.L., Paternoster, I.L., Singh, H., Storer, R., Weingarten, G.G., Williamson, C., Brodie, A.C., Cook, J., Lake, D.E.,. Meerholz, C.A., Turnbull, P.J., and Highcock, R.M. J Chem Soc Perkin Trans I, 1991; 2467. Komatsu, H. Araki, T. Tetrahedron Lett. 2003; 44: 2899. Herdewijn, P., Balzarini, J., Baba, M., Pauwels, R., Van Aerschot, A., Janssen, G., De Clercq, E. J Med Chem. 1988; 31: 2040. Komatsu, H., Awano, H., Ishibashi, H., Ikeda, I. Nucleic Acid Research, 2001, suppl. 1: 49. Holguin-Hueso, J. Cardinaud, R. FEBS Lett. 1972; 20: 171. Pochet, S., Dugué, L., Meier, A., and Marliere, P. Bioorg Med Chem Lett. 1995; 5: 1679.
Bioprocesses for the Synthesis of Nucleosides and Nucleotides
93
[70] Okuyama, K., Shibuya, S., Hamamoto, T., and Noguchi, T. Biosci Biotechnol Biochem. 2003; 67: 989. [71] Carson, D.A. and Wasson, D.B. Biochem Biophys Res Commun. 1988; 155: 829. [72] Bavaro, T. Scientifica Acta, 2007; 1: 65. [73] Ferrero, M. and Gotor, V. Chem Rev. 2000; 100: 4319. [74] Rocchietti, S., Terreni, M. and Pregnolato, M. 2003 WO03057894. [75] Uemura, A., Nozaki, K., Yamashita, J., and Yasumoto, M. Tetrahedron Lett. 1989 30: 3819 [76] Zinni, M.A., Rodríguez, S.D., Pontiggia, R.M., Montserrat, J.M., Iglesias, L.E., and Iribarren, A.M. J Mol Cat B:Enzymatic, 2004; 29: 129. [77] Singh, H.S., Cote, G.L. and Sikorski, R.S. Tetrahedron Lett 1993; 34: 5201 [78] Panero, J., Trelles, J., Rodano, V., Montserrat, J.M., Iglesias, L.E., Lewkowicz, E.S., and Iribarren, A.M. Biotechnol. Lett. 2006; 28: 1077. [79] Pregnolato, M., Terreni, M., Ubiali, D., and Bavaro, T. 2008; PCT/IB2008/000482. [80] Capello, M., Gonzàlez, M., Rodríguez, S.D., Iglesias, L.E., and Iribarren, A.M. J Mol Cat B:Enzym. 2005; 36: 36. [81] Capello, M., Imanishi, L., Iglesias, L.E., and Iribarren, A.M. Biotechnol. Lett. 2007; 29: 1217. [82] Damkjaer, D.L., Petersen, M. and Wengel, J. Nucleosides Nucleotides, 1994; 13: 1801. [83] Sugira, M. and Isobe, M. Chem. Phar. Bull. 1976; 24: 1822. [84] Brzozowski, A.M., Derewenda, U., Derewenda, Z.S., Dodson, G.G., Lawson, D.M., Turkemburg, J.P., Bjorkling, F., Huge-Jensen, B., Patkar, S.S., and Thim, L. Nature, 1991; 351: 491; b) Derewenda, U., Brozowski, A.M., Lawson, D.M., and Derewenda, Z.S. Biochemistry, 1992; 31: 1532; c) Derewenda, Z.S. and Derewenda, U. J Mol Biol. 1992; 227: 818; d) Derewenda, Z.S. Structural biology, 1995; 2: 347; e) Grochulski, P., Li, Y., Schragm, J.D., Boutthilier, F., Smith, P., Harrinson, D., Rubin, B., and Cyler, M. J Biol Chem. 1993; 268: 12843. [85] Fernández-Lafuente, R., Armisen, P., Sabuquillo, P., Fernández-Lorente, G., and Guisán,, .JM. Chem Phys Lipids, 1998; 93: 185. [86] Fernandez-Lorente, G., Terreni, M., Mateo, C., Bastida, A., Fernández-Lafuente, R., Dalmases, P., Huguet, J., and Guisán, J.M. Enzyme Microb Technol. 2001; 28: 389; b) Palomo, J.M., Fernandez-Lorente, G., Mateo, C., Fuentes, M., Fernandez-Lafuente, R., and Guisan, J.M. Tetrahedron: Asymmetry, 2002; 13: 1337; c) Palomo, J.M., Filice, M., Fernandez-Lafuente, R., Terreni, M., and Guisan, J.M. Adv Synth Catal. 2007; 349: 1969; d), Fernandez-Lorente, G., Filice, M., Terreni, M., Guisan, J.M., FernandezLafuente R, and Palomo J.M. J Mol Cat B:Enzymatic, 2008; 51: 110. [87] Bastida, A., Sabuquillo, P., Armasen, P., Fernández-Lafuente, R., Huguet, J., and Guisán, J.M. Biotechnol Bioeng. 1998; 58: 486. [88] Fernandez-Lorente, G., Palomo, J.M., Coca, J., Mateo, C., Moro, P., Terreni, M., Fernandez-Lafuente, R., and Guisan, J.M. Tetrahedron. 2003; 59: 5705. [89] Kuninaka, A. and Shoyu, Y. Hydrolysis of nucleic acids with Penicillium nuclease. In: Microbial Production of Nucleic Acid-Related Substances. Ogata, K., Kinoshita, S., Tsunoda, T., and Aida, K., Eds. (Tokyo: Kodansha Ltd), 1976 pp. 75–86. [90] Matsui, H., Sato, K., Enei, H., and Takinami, K. Agric Biol Chem. 1982; 46: 2347.
94
Marco Terreni, Daniela Ubiali, Teodora Bavaro, et al.
[91] Matsui, H., Sato, K., Enei, H., and Hirose, Y. Agric Biol Chem. 1979; 43: 1739. [92] Arner, E.S.J. and Eriksson, S. Pharmacol Ther. 1995; 67: 155. [93] Bradshaw, H.D., Jr. and Deininger, P.L. Mol Cell Biol. 1984; 4: 2316; b) Chottiner, E.G., Shewach, D.S., Datta, N.S., Ashcraft, E., Gribbin, D., Ginsburg, D., Fox, I.H., and Mitchell, B.S. Proc Natl Acad Sci 1991; 88: 1531; c) Johansson, M. and Karlsson, A. Proc Natl Acad Sci. 1996; 93: 7258; d) Wang, L., Hellman, U. and Eriksson, S. FEBS Lett. 1996; 390: 39; e) Johansson, M. and Karlsson, A. J Biol Chem. 1997; 272: 8454; f) Wang, L., Munch-Petersen, B., Herrström-Sjöberg, A., Hellman, U., Bergman, T., Jörnvall, H., and Eriksson, S. FEBS Lett. 1999; 443: 170. [94] Verri, A., Montecucco, A., Gosselin, G., Boudou, V., Imbach, J.L., Spadari, S., and Focher, F. Biochem J. 1999; 337: 585. [95] Munch-Petersen, B., Piskur, J. and Sondergaard, L. J Biol Chem. 1998; 273: 3926. [96] Ihlenfeldt, H.G., Munch-Petersen, B., Piskur, J., and Sondergaard, L. 2006; US patent 2006252131. [97] Bastner Sandrini, M.P., Söderbom, F., Mikkelsen, N.E., and Piskur, J. J Mol Biol. 2007; 369: 653. [98] Montgomery, J.A. and Shortnacy, A.T. 1982; U.S. Patent 4,357,324. [99] Liberman, J., Korncberg, A. and Simms, E. J Am Chem Soc. 1954; 76: 3608; b) Florid, A., Palmarini, C., Fini, C. J Chromatogr. 1977; 138: 212. [100] Bochkov, D.V., Khomov, V.V. and Tolstikova, TG. Biochemistry, (Moscow) 2006; 71: 79. [101] Barai, V.N., Kvach, S.V. and Zinchenko, A.I. Mikhailopulo IA. Biotechnology Letters, 2004; 26: 1847. [102] a) Mihara, Y., Utagawa, T., Yamada, H., and Asano, Y. 2000 US 006,015,697A; b, Mihara Y, Utagawa T, Yamada H, Asano Y. Appl Environ Microb. 2000; 66: 2811. [103] Rossolini, G.M., Schippa, S., Riccio, M.L., Berlutti, F., Macaskie, L.E., and Thaller, M.C. Cell Mol Life Sci. 1998; 54: 833. [104] Kier, L.D., Weppelman, R. and Ames, B.N. J Bacteriol. 1977; 130: 399.
In: Biocatalysis Research Progress Editors: F. H. Romano, A. Russo
ISBN: 978-1-60456-619-2 © 2008 Nova Science Publishers, Inc.
Chapter III
Biocatalysis in Environmental Technology D. M. G. Freire1,*, M. L. E. Gutarra1, V. S. Ferreira-Leitão2, M. A. Z. Coelho3 and M. C. Cammarota3,† 1
Departamento de Bioquímica, Instituto de Química, Universidade Federal do Rio de Janeiro, Av. Athos da Silveira Ramos, 149, Centro de Tecnologia, Bloco A, Lab 549 (1 e 2), Cidade Universitária, Ilha do Fundão, CEP: 21941-909, Rio de Janeiro, R.J., Brazil 2 Divisão Ambiental - Instituto Nacional de Tecnología, Av. Venezuela, 82, Sala 302, Centro, Rio de Janeiro, R.J., CEP: 20081-312, Brazil 3 Departamento de Engenharia Bioquímica, Escola de Química, Universidade Federal do Rio de Janeiro, Av. Horácio Macedo, 2030, Centro de Tecnologia, Bloco E , Sala 203, Cidade Universitária, Ilha do Fundão, CEP: 21941-909, Rio de Janeiro, R.J., Brazil
Abstract New strategies development to solve problems related to waste disposal may include technologies facing compounds of low biodegradability and/or detoxification of some residues with the alternative of adding value. There are various chemical and biological approaches, but biological ones employing enzymes had been showed good results. On the other hand, environmental use of enzymes depends on costs reduction for biocatalyst production through the selection of highly-productive strains, development and optimization of fermentation processes. Production system which employs agroindustrial residues as culture medium can be an interesting alternative. Microorganisms’growth on solids wastes by solid state fermentation can add value to these residues by enhancing
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D. M. G. Freire, M. L. E. Gutarra, V. S. Ferreira-Leitão et al. nutritional quality and/or producing several enzymes with biotechnological applications, and also can reduce its pollution potential by degrading toxic compounds. The reduction cost of enzyme production could make feasible its employment for environmental purposes such as enzymatic pretreatment of wastewaters with high levels of fats, improving the biological degradation, or in the treatment of colored and phenolic wastes. The potential of environmental biocatalysis not only help to solve pollution problems but also simultaneously add value to undesirable wastes by the generation of biotechnological products or by enhancing its nutritional proprieties for use as animal feed.
1. Introduction The sustainable development promotion is reached when the productive sector acts in an integrated way with the environment. This proposal, one of the green chemistry pillars, gives redirection of the traditional industrial model, in order to obtain a minimal emission. This situation is achieved when an industry makes use of its own residue as crude material, or another industry can use it to carry out a new cycle of production: industries organizing themselves in clusters, in a way that residues or byproducts without value in an activity can be converted in an aggregated value input to the others. This paradigm change puts in question a plenty of industrial production technologies that do not take into account, adequately, the environment maintenance. It is necessary to plan and restructure the industrial production in order to have all the raw material transformed in useful property or reintegrated in the ecosystem without damage it. This procedure foresee plenty of opportunities for research and new technologies development to maximize the raw material use to complete the material cycle or the residues discarded in the ecosystems. In this context, the development of new strategies to solve problems related to waste disposal may include technologies driven to compounds of low biodegradability and/or detoxification of some residues providing value addition alternatives. Although there are different chemical and biological approaches, the future intensive application of enzymes in environmental purposes depends on cost reduction of biocatalyst production through the selection of highly-productive strains, development and optimization of fermentation processes. On the other hand, the biocatalyst production employing agro-industrial residues can be an interesting alternative to solve pollution problems. The solid state fermentation with filamentous fungi, yeast or bacteria can also add value to these residues by enhancing nutritional quality or producing several enzymes with biotechnological application. For instance, attempts have been made to detoxify residues through biological means as in solid state fermentation by fungus Penicillium sp.. The fungus has eliminated toxic content to no detectable level, reduced allergenic potential and produced simultaneously high contents of hydrolases. Enzyme produced at low cost by solid state fermentation can be employed in enzymatic pretreatment of wastewaters from dairies, slaughterhouses and meat processing plants with high levels of fats and proteins to improve its biological degradation. Such strategy has been previously adopted enabling a considerable increase in organic matter removal efficiency and resulting in the attainment of a high-quality
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effluent. The possible employment of the hydrolyzed fat to increase biogas production in posterior anaerobic process is a goal of such approach. In the last decades, the use of enzymes and specific microorganisms in the treatment of colored and phenolic wastes has also been object of several scientific works. Oxidative enzymes (laccases, peroxidases and polyphenol oxidases) produced by white-rot fungi or plant residues extracts catalyze oxidative and nucleophilic reactions leading to unstable products which precipitate, being thus removed from the aqueous solution. Both microbial and enzymatic treatments are efficient for decolorization and the choice for industrial applications may consider economic and safety aspects. The potential of environmental biocatalysis not only helps to solve pollution problems but also leads to simultaneous production of biotechnological compounds with added values.
2. Biocatalysts Production for Environmental Technology Application The main used procedures in enzyme production are named submerged fermentation (SmF) and solid state fermentation (SSF). Nearly 90% of all industrial enzymes are produced by submerged fermentation, frequently using genetically modified microorganisms (Hölker and Lenz, 2005). However, in environmental applications the cost and the use of genetically modified organisms (GMOs) can make unfeasible the later usage of these enzymes in the environment (Cammarota and Freire, 2006). SSF was reported by Chineses in 1000 a.C. as the used technique to obtain the soy sauce. So, SSF is an employed practice in East, since centuries ago, to obtain fermented beverage and food, such as: “chiang”, “sufu” and “red rice” in China; “tempe” and “ontjom” an Indonesia; “miso”, “hamanatto”, “natto” and “sake” in Japan (Mitchell et al., 2002). This technology was introduced in West around 1890 when a Japanese named Takamine established, in the United States of America, an enzyme industry based in this oriental technology. Historically, the submerged fermentation (SmF) development in stirred tanks, due to antibiotic industry development around II World War period, had as consequence a meaningful decrease in the released attention to the SSF process. As consequence, in West there was an expansion of equipments and production monitoring technology development seeking SmF. In Japan and other East countries, mainly due to their food culture that includes fermented foods, it preceded emphasizing SSF (Hesseltine, 1972). The main difference between these fermentative processes is the free water amount into culture medium. In SmF, solid amount does not surpass 50 g.L-1, whereas in SSF the solid contents can vary from 20 to 70% of total weight. Thus, SSF is a process that occurs in absence or near-absence of free water. (Mitchell et al., 2002). SSF can be defined as a process that employes natural substrates that act as carbon and nitrogen source (Pandey et al., 2000; Mitchell and Lonsane, 1992). The main differences between SSF and SmF are presented in Table 1 (Mitchell and Lonsane, 1992).
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Solid State Fermentation (SSF) Culture medium without liquid phase Water insoluble substrates Water is in the medium in enough amount for the microorganism growth The nutrient absorption occurs from the moistened solid substrate Gradients presence of heat, nutrients, products and O2 Presents three phases: solid, gaseous and liquid Discontinuous liquid phase Greater inoculum concentrations Aeration makes available O2 and removes the metabolic heat and gaseous product Facultative agitation More concentrated products The fungic growth comprehends hyphae penetration into the substrates Bacteria and yeast grow adhered to the solid substrate
Submerged Fermentation (SmF) Culture medium with liquid phase Water soluble substrates A lot of water The nutrient absorption occurs from the dissolved nutrients Heat, nutrients, products and O2 are evenly scattered Presents two phases: liquid and gaseous Continuous liquid phase Lower inoculum concentrations Aeration makes available O2 and gaseous product, but the metabolic heat is removed by peculiar apparatus for heat changes Agitation is generally essential Less concentrated products The fungic growth occurs in individual form of mycelium or aggregations evenly issued in the stirred medium Bacteria’s and yeast grow evenly distributed into the stirred medium
Solid state fermentation (SSF) is becoming a good alternative for enzyme production, aiming at environmental application, with low cost, from wild microorganisms. This fermentation type presents as advantage to enable the use of its own fermented medium as enzyme source, generally an agro industrial solid residue. SSF features grant it several advantages over submerged fermentation, mainly when the final target is the production of biocatalysts at low cost to be used in environmental technology. One of the main advantages of SSF is the use of simple culture medium, from vegetable origin material, such as bran and rice peel, wheat, corn and other cereals, demanding a little amount of additional nutrients into the medium. This advantage provides the use of agro industrial residues as substrate, representing, in several countries, abundant and low cost raw material. In SSF, a little amount of water is employed, implying in several advantages, as lower content of reaction medium; lower area demanded for bioreactor related to the product yield; and higher product concentration. Moreover, less effluent amount is also generated in the process; it is not necessary the addition of anti-foaming agents; and the contamination by bacterias and yeast is lower due to the lower water activity in the medium (Hölker and Lenz, 2005). Furthermore, the enzyme production in SSF by filamentous fungi shows, many times, advantages over SmF production, respecting enzyme stability and productivity (Tsuchiya et al., 1994; Mateos Diaz et al., 2006) or product obtainment with more interesting characteristics (Acuña-Arguelles et al., 1995).
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Hydrolisis and esterifications O
+
R1 O
R2
H2O
O
enzyme
R1 OH
+ R2
OH
Transesterifications a) acidolysis O R1 O
O
O R3
+
R2
enzyme
O
R3 O
OH
+
R2
R1 OH
b) alcoholysis O
O
R1 O
R2
+
enzyme
R3
R1 O
OH
R3
+
R2 OH
c) interesterification
R1 O
O
O
O R2
+
enzyme
R3 O
O
R1 O
R2
R4
+
R3 O
R2
d) aminolysis O
O
enzyme
R1 O
R2
+
R3
NH2
R1 NH
R3
+ R2
OH
Figure 1. Reactions catalyzed by lipases.
Another advantage is that SSF presents culture conditions nearer microorganisms natural habitat conditions, mainly the filamentous fungi, allowing the obtainment of the product in a great deal. (Rivera-Muñoz et al., 1991; Murthy et al., 1993; Nigam and Singh, 1994). However, there is not an established scale or a method that allows a direct comparison of the productivity between these two systems in actual terms (Pandey, 2003; Viniegra-González et al., 2003; Azeredo et al., 2007). Another aspect, still controversy, raised in literature as this fermentation advantage is related to its capacity in supplanting the catabolic effects by easily assimilable substrates. Several hypothesis have been proposed to explain differences in enzyme activity titers between SmF and SSF, among which are the water content, the nature of SSF, the diffusion of nutrients on solid matrix and the changes in the ratio between the substrate uptake rate and diffusivity substrate coefficient (Aranda et al., 2006). In SSF, the lower mass transfer processes, related to gases and nutrients diffusion, are strongly influenced by the physical structure of the matrix and by the liquid phase of the system, and are limited to diffusion (Papagianni et al., 2001; Pérez-Guerra et al., 2003). Recently, Cerda-Montalvo et al. (2005) demonstrated, experimentally, the effect of substrate diffusion on enzyme expression. Nutrients diffusion occurs at an intraparticular level and includes both the diffusion of nutrients toward the cells and the hydrolysis of solid substrates by the microbial enzymes. This last point is an important aspect in SSF because most of the substrate is water insoluble.
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In contrast, microorganisms in SmF are exposed to hydrodynamic forces, and consequently better mass transfer and higher nutrient availability are present (Papagianni et al, 2000 and 2001). Although countless SSF advantages over SmF, SSF is a heterogeneous system, where the nutrients and oxygen arrangement, the product concentration and the growth are not well distributed through the entire medium and the stir utilization requires high energy consumption. Beyond that, the medium heterogeneity implies in difficulties on monitoring and parameters control, very important for the fermentative process, as temperature, pH, moisture and oxygen. These various technical difficulties with large-scale solid-state cultivation processes mean that submerged liquid cultivation, for which the technology developed significantly over the last half of the twentieth century, is still the dominant cultivation method for biotechnological applications. However, in certain situations solidstate cultivation gives advantages over submerged liquid cultivation, especially in its various environmentally-related applications.
3. Main Biocatalysts Employed in Environmental Technology 3.1. Lipases Lipases (triacylglycerol ester hydrolase E.C. 3.1.1.3) and esterases (E.C. 3.1.) are biocatalysts that have an extensive and diversified application field, reason for which their participation in the industrial enzyme worldwide market is increasing meaningfully. It is estimated that, in the future, they will present so high significance as proteases have nowadays. (Sharma et al., 2001; Cammarota and Freire, 2006). These enzymes constitute a wide family that have as natural catalytic function the hydrolysis of triacylglycerol ester bounds, releasing diglycerides, monoglycerides, glycerol and free fatty acids. However, under non-aqueous conditions can occur the reverse reaction (esterification) or also several transesterification reactions (Figure 1). Lipases occur widely in nature, produced by many microorganisms and higher eukaryotes. In animals, lipases obtained from pig and human pancreas are best known and most investigated than all other lipases. In these organisms they are engaged in several lipid metabolism steps, including fat digestion, adsorption, reconstitution and lipoproteins metabolism. In plants, lipases are present in higher plants seeds, as castor bean (Ricinus communis) and canola (Brassica napus), they are also found in several plants energy reserve tissues (Jaeger and Eggert, 2002; Sharma et al., 2001; Villeneuve, 2003; Cavalcanti et al., 2007). However, for enzymes industrial production, microorganisms are the preferred source, once they have shortest generation time, high yield of substrate conversion product, great versatility and major simplicity in genetic manipulation and in cultivation conditions. Due to habitats multiplicity, microorganisms produce distinct lipases types, regarding specificity to certain substrates and also in optimum pH and temperature range. Lipases are produced by bacterias, filamentous fungi and yeast, allowing these microorganisms to use lipids from animal or vegetable origin as carbon and energy sources for their growth. Though many
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microorganisms have been reported in literature as lipase producers, the genera Candida, Rhizopus and Pseudomonas are considered the main lipase industrial sources. The yeast Candida rugosa is the most employed microorganism for lipase production. There are commercial products obtained from this yeast being trading by several companies as Amano, BDH, Biocatalysts, Roche, Fluka, Meyto-Sangyo e Sigma (Ferrer et al., 2002). Though, aiming the productivity increase of enzyme production by these microorganisms, genetic engineering has achieved several important advances (Bornscheuer et al., 2002). However, a microbial wild strain able to produce high levels of enzyme presents advantages related to major agent stability and to more enzyme approval facility for use, for instance, in food industry and in the environment. In this context, the great biodiversity of tropical countries represents a source practically inexhaustible in the search for wild species lipase producers in a great deal and/or with new properties. There are several works in literature about microbial lipase production in SmF as well as in SSF. In submerged fermentation several works are being accomplished aiming fix the optimum medium cultivation composition, identifying the concentration influence and the origin of carbon and nitrogen sources used for lipase production (Sharma et al., 2001). Lipids addition is, generally, essential for high lipase activities obtainment in this system type (Pokorny et al., 1994; Freire et al., 1997). The effects of different concentrations of perfluorodecalin (PFC), used as an oxygen vector in SmF to enhance lipase productivity by increasing the oxygen availability to Yarrowia lipolytica, has been investigated by Amaral et al. (2006) leading the 23-fold enhacement with the addition of 20% (v/v) PFC. Moreover, it is shown that using PFC and glucose as substrate is more effective in lipase production than the conventional use of olive oil. Dalmau et al. (2000) and Azeredo et al. (2007), have studied lipase production from Candida rugosa and Penicillium restrictum, respectively, in SmF and observed that lipids and fatty acids utilization as carbon sources allows greater yields and glucose presence provoked a lipase production repression effect. This catabolite repression phenomenon is oftentimes observed in SmF (Freire et al., 1997, Dalmau et al., 2000), but in SSF it seems to be reduced (Ramesh e Lonsane, 1991), due to, mainly, the mass transfer inherent to the system (Azeredo et al., 2007). In SSF, lipases are being produced in several solid residues, as wheat meal (Mahadik et al., 2002; Sun and Xu, 2008), sesame oil cake (Kamini et al., 1998), rice meal (Rao et al., 1993), soy meal (Kempka et al., 2007); physic nut cake (Mahanta et al., 2008) and babassu cake (Gombert et al., 1999; Palma et al., 2000; Gutarra et al., 2005 and 2007). The solid residues employed as culture medium for SSF, can be supplemented with different carbon and nitrogen sources. Lipids are frequently employed as lipase inducers; however the precise mechanism leading to enhancement of enzyme production by lipid compounds, which can hardly penetrate into the cell, is yet to be elucidated. Carbohydrates supplementation can also allow high activities lipase obtainment (Gombert et al., 1999; Azeredo et al., 2007). Gombert et al. (1999) studied the lipase production by Penicillium restrictum in SSF, using babassu cake supplemented with peptone, olive oil and Tween-80 as culture medium. All employed supplements provoked a lipase production increase, and peptone supplemented showed the best results. In this work, it was observed that in lipase production, the supplement type practises an influence grater than the ratio C:N. Lipase production obtained by Penicillium verrucosum using soy bean meal as substrate was investigated by Kempka et al. (2007).
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Different inductors were evaluated and the results showed that there is no influence of this variable on the lipase production, while temperature and initial moisture were the main factors that affected this enzyme production. The optimized cultivation temperature (27.5 oC) and initial moisture of substrate (55%) resulted in lipase activities of 40 U/g of dry meal. Although the effects of media composition in lipase production have been extensively investigated, the physiological mechanisms, which govern the microbial response to change in the environmental conditions, remain unclear and should be better understood (Bornscheuer et al., 2002; Gombert et al., 1999). Several other works compared lipase production in SSF and SmF, all of them report greater enzyme production and stability when SSF system was employed. Azeredo et al. (2007) observed completely different physiological behaviors after the addition of easily (oleic acid and glucose) and complex (olive oil and starch) assimilable C sources for the liquid and solid media (babassu cake). High lipase production using glucose was only attained in SSF, maybe due to such fermentation ability in minimizing catabolite repression. In opposition to these results, Gutarra et al. (2005), working with another filamentous fungus (Penicillium simplicissimum) in the same residue (babassu cake) did not observe significant differences related to easily and complex carbon sources in both fermentation types. So, this SSF characteristic of turn around the catabolic repression by easily assimilation carbon sources seems to be dependent of the substrate type and microorganism. Beyond the technical aspects, economic issues must also be taken into account when considering the use of enzymes in environmental technology. In this context, the use of solidstate cultivation to produce enzymes to environmental application has proved to be more economical than the use of submerged cultivation techniques. Using Penicillium restrictum and babassu oil cake for the production of a lipolytic “liquid enzyme product”, considering a production scale of 100 m3 per year, the total capital investment needed for the solid-state cultivation process is 78% lower than that needed for the submerged process and the unitary production cost for solid-state cultivation is 47% lower than the market price of an existing liquid lipolytic product. The solid-state cultivation process is very attractive from an economic point of view, with a payback time of 1.5 years, a return on investment of 68% and an internal return rate of 62% for a 5-year-project life (Castilho et al., 2000). These economic advantages of solid-state cultivation for the production of hydrolytic enzymes are mainly due to the low capital investment and to the very cheap raw material used.
3.2. Proteases Proteases represent one of the three greatest enzyme groups more important for industry, accounting for about 60% of enzymes worldwide sell, where 40% of these enzymes are from microbial origin (Rao et al., 1998). Whereas enzymes are a physiologically requisite to living being, they are found in wide source diversity, such as plants, animals and microorganisms. The increasing interest for microbial enzyme commercially produced is due to the shortage ability that proteases from vegetable or animal origin have in attending the worldwide demand. Proteases are a single class of enzymes that catalyze the proteins hydrolysis through peptides bond cleavage. According to enzyme nomenclature, proteases are classified in
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subgroup 4 of group 3 of hydrolases. However, due to the huge action and structure diversity, proteases do not follow easily the classification system and the nomenclature created for enzymes in general. Such fact has generated efforts for the development of other classification shape for this specific group. Three criteria are in use for peptidase classification: the catalyzed reaction; the chemical nature of the catalytic site; and the evolutionary relationship revealed by its structure. Proteases are roughly subdivided in two main groups, depending on its action site: exopeptidases (amine and carboxypeptidases) and endopeptidases (serine, aspartic, metallo and cysteine proteinase) (Barrett, 1994). Proteolytic enzymes production by different microorganisms using submerged fermentation process (SmF) has been widely described (Yang and Wang, 1999; Singh et al., 2001; Gessesse et al., 2003; Thys et al., 2004). In SSF production, filamentous fungi can be quoted as the most commonly employed microorganism in this fermentation type, mainly due to their good growth in these low moisture conditions (Malathi and Chakaborty, 1991; Germano et al., 2003). Attempts for SSF enzyme production using bacteria have also been done successfully in the last years (Pandey et al., 2000; Uyar and Baysal, 2003; Soares et al., 2005; Mahanta et al., 2008). The microbial production of thermophilic and keratin hydrolyzing proteases by a streptomycete using a low cost medium, composed by industrial poultry and corn processing by-products was described by Lima et al. (2006). Protease production by Streptomyces sp. 594 was obtained by submerged (SmF) and solid state fermentation (SSF) using feather meal (FM) and corn steep liquor (CSL) as sole sources of carbon (C) and nitrogen (N). These proteases, which belong to serine and metalloproteinase classes, were active at high temperatures (55 to 90 °C), and over a wide range of pH values (5.0 to 10.0). This thermophilic and keratin hydrolyzing proteases have shown interesting properties for industrial purposes (Azeredo et al., 2006).
3.3. Phytases and Cyanidases Phytases (E.C.3.1.3.26) are meso-inositol hexaphosphate phosphohydrolases that catalyze the stepwise phosphate splitting of phytic acid (IP6) or phytate to lower inositol phosphate esters (IP5-IP1) and inorganic phosphate (Lei and Porres, 2003). Figure 2 illustrates a phytase action scheme in its natural substrate - phytate. This enzyme used in several feed sort has fundamental importance, because allows the increasing bioavailability of several nutrients. In many raw materials from vegetable origin used in animal feed (cereal grains, legumes and oleaginous seeds), a great deal of mineral phosphorus (75-80%) is storaged in phytic acid form (myo-inositol 1,2,3,4,5,6 – hexakisphosphate or phytate), that is not degraded by monogastric animals, like pigs, chickens and fishes (Pandey et al., 2001). Besides perform phosphorus unavailable, phytate binds itself with divalent cations, such as Ca2+, Mg2+, Zn2+, Cu2+, Mn2+and Fe2+, avoiding these nutrients absorption in the animal bowels. Thus, phytases besides have hand in a better digestion of certain components in monogastric animals (phosphorus utilization), also contribute for decrease the levels of phosphorus excreted by the animal, with following pollution reduction (Lei and Porres, 2003).
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Figure 2. Phytase action mode.
A number of phytase genes and proteins have been identified from plants and microorganisms including bacteria, yeast, and filamentous fungi and spite of yeast and bacteria production ability, the last ones are more employed (Pandey et al., 2001). The first and better characterized commercial phytase products derived from Aspergillus niger, with the capacity to release phytate-bound phosphorus and reduce phosphorus excretion, was introduced into market in 1991 (Selle and Ravindran, 2007). Phytase production by Aspergillus fungus genus has been extensively studied by several authors through SmF as well as SSF (Kim et al., 1999; Papagianni et al., 2000; Ebune et al., 1995). Several other filamentous fungi are well known to produce phytases in SSF (Pandey et al., 2001; Bogar et al., 2003). Thermophilic fungi have not been explored adequately like their mesophilic counterparts. Chadha et al. (2004) optimized phytase production by a thermophilic mould Rhizomucor pusillus in SSF using wheat meal as substrate. The fungal SSF product contains not only phytase, but also accessory enzymes, fungal protein and organic acids which increase feed digestibility by permitting the access to phytates present in it. Sporotrichum thermophile has recently been shown to produce phytase in SSF using sesame oil cake as the substrate and its suitability in the dephytinization of sesame oil cake. This mould also secreted phytase in cane molasses medium and its production was optimized in submerged fermentation (Singh and Satyanarayana, 2006). Bogar et al. (2003) reported phytase production by A. ficuum NRRL3135, Mucor racemosus NRRL1994 and Rhizopus oligosporus NRRL 5905 on canola meal, cracked corn, soybean meal and wheat meal in SSF. Optimization studies were carried out using Plackett-Burman and central composite designs showing the improved production when wheat meal was supplemented with starch and ammonium sulphate. The phosphorous concentration effect has been studied in phytase
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production and several works indicate that high phosphate conditions are known to repress the synthesis of acid phosphatases and phytases, while limiting phosphate conditions result in their expression (Vats and Banerjee, 2004). Another enzyme very useful in detoxification of cassava roots, for feed application, is cyanidase or linamarase. Linamarase is a cyanogenic ß-glucosidases, which catalyses the hydrolysis of linamarin into glucose and acetone cyanohydrin (Giraud et al., 1992). Cyanides function as a defense mechanism against attack by predators and are distributed mostly in leaves and roots cortex (peel) (in higher quantities) and in smaller quantities in the parenchyma (interior) (Cardoso et al., 2005).
3.4. Tannases Tannin acyl-hydrolase (E.C. 3.1.20), or tannase, catalyzes the hydrolysis of ester bonds in gallotannins, complex tannins and gallic acid esters. This enzyme is employed in food and beverage processing; however, it’s practical use is limited in consequence of insufficient knowledge about its properties, expression, and large-scale application (Aguilar et al., 2007). Figure 3 illustrates tannase action in its natural substrate - tannins. Despite being present in plants and animals, it has been mainly produced by bacteria, yeast and fungus. Filamentous fungi like Aspergillus and Penicillium genera and bacteria as Bacillus and Lactobacillus genera have been widely used in tannase production (Mondal et al., 2001; Pinto et al., 2001; Deschamps et al., 1983; Bradoo et al., 1996; Batra and Saxena, 2005; Nishiatani et al., 2004; Kostinek et al., 2007). Not all tannases are equally active against the different tannin substrates. Fungal tannases have a better activity in degrading hydrolysable tannins, whereas yeast tannases are better in tannic acid degradation and have a lower affinity for naturally occurring tannins. On the other hand, bacterial tannase can degrade and hydrolyze natural tannin and tannic acid very efficiently (Deschamps et al., 1983). Of all the microorganisms able to produce tannase, Aspergillus sp. was commercially the most efficient producers of this enzyme. Studies on tannase production by Aspergillus have been carried out in submerged and solid-state cultures. Depending on the strain and the culture conditions, the enzyme is induced and expressed with different levels of activity, showing distinct production patterns. However, the induction mechanism has not been clearly demonstrated, and there are some controversies about the role of some of the hydrolysable tannin constituents as related to the synthesis of tannase (Deschamps et al., 1983; Aguilar et al., 2001). In spite of tannase production have been strongly studied in submerged fermentation, several authors have been researching this enzyme production using agro industrial residues, such as wheat meal; tamarind seed powder; palm kern cake (Sabu et al., 2005) and coffee rusk blend to wheat meal (Battestin and Macedo, 2007). The principal advantages related to solid medium cultivation are higher productivity (Aguilar et al., 2002) and higher pH and temperature stability (Lekha and Lonsane, 1994; Rana and Bhat, 2005) in achieved preparations by solid state fermentation.
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Figure 3. Tannic acid Hydrolysis.
Figure 4. Reaction scheme of tyrosinase.
3.5. Polyphenol Oxidases Polyphenol oxidase or tyrosinase (EC 1.14.18.1) is an oxygenase oxyreductase. This enzyme has been found widely distributed throughout the phylogenetic scale from bacteria to mammals including the mushroom Agaricus bisporus (Van Gelder et al., 1997). In this fungal structure, the enzyme is readily available leading to high activity extracts without extensive purification. This suggests its potential as a biocatalyst for applications involving biomodification of phenols or bioremediation of phenol-polluted water (Kameda et al., 2006). Tyrosinase catalyzes two different reactions that occur in series: the hydroxylation of monophenol to o-diphenols (monophenolase or cresolase activity) and the oxidation of odiphenols to o-quinones (diphenolase or catecholase activity), both using molecular oxygen (Van Gelder et al., 1997) (Figure 4). These quinones are unstable in aqueous solutions and perform a non-enzymatic polymerization through oxidative and nucleophilic reactions. The product of such polymerization presents dark color and according to the reaction substrate precipitates after some hours. (Duckworth and Coleman, 1970; Atlow et al., 1983; Wada et al., 1993). Besides, tyrosinase could be inactivated as a result of irreversible reaction between
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a product of enzyme-catalyzed reaction and the active site of the enzyme, called suicide inactivation by Escribano et al. (1989). Peroxidase is known to be capable in removing phenolic groups and aromatic amines in aqueous solutions and also performs decolorization of textile wastewater (Nakamoto and Machida, 1992; Tong et al., 1998). But this enzyme needs hydrogen peroxide as an oxidative agent while tyrosinase uses molecular oxygen leading to a less expensive process. Both enzymatic treatments (with peroxidase or tyrosinase) provide the removal of phenolic compounds from industrial wastewater. Since most dyes usually employed in textile industry presents phenolic groups in its chemical structure (Figure 5), the tyrosinase potential in decolorizing textile effluents through the action of this enzyme (polymerization and dye precipitation) is being evaluated. Most works reported in the literature use the enzyme extract from mushrooms (Agaricus bispora) according to the methodology described in Atlow et al. (1983). For instance, different values of enzymatic activity were observed. Cammarota and Coelho, 2006) obtained a mean activity in the crude extract of around 511 U/mL and a specific activity of 852 U/mg. These values are lower than those achieved by Atlow et al. (1983) employing the same protocol: 2800 U/mL. Tyrosinase purchased from Sigma-Aldrich presents a specific activity between 2800 - 3500 U/mg. Such large activity variation was investigated by Bevilaqua (2000), verifying that the tyrosinase activity varies with the mushrooms maturation grade, aging after-harvest and freezing time of the mushrooms cake. The influence of the mushroom age in the enzyme activity is uncertain, acting in latent forms of the enzyme or causing an increase in enzyme inactivation due to natural cell lysis. Longer freezing times lead higher formation of ice crystals that enhance tyrosinase extraction from cell disrupt. Although, polyphenol oxidase could be produced by others sources, the worldwide market of mushroom has increased and consequently the costs of enzyme extraction decreased.
Figure 5. Chemical structure of a reactive dye, Reactive Black 5 (Remazol Black GF).
3.6. Laccases and Peroxidases Ligninolytic enzymes have a great potential for industrial application and for this reason, many groups are devoted to their characteristics, properties and applications studies (Couto and Herrera, 2006; Couto and Sanroman, 2005; Wesemberg, 2003; Duran et al., 2002; Duran and Espósito, 2000). These enzymes have as main function in nature the lignin degradation, a natural non-hydrolysable biopolymer that constitutes the greatest natural source of aromatic
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structures (Bietti et al., 1998; Reid, 1995; Tuor et al., 1995). White rot fungi are the best known and studied producers, despite of several works report yeast and bacteria ability in producing ligninolytic enzyme complex (Yang 2008; Gottschalk et al., 1999; HernandezPérez et al., 1998). The principal involved enzymes in this complex are peroxidases (lignin peroxidase and manganese peroxidase) and laccases. This natural oxidation ability of a complex and recalcitrant structure confers to ligninolytic enzymes very wide and diversified properties, which can be applied from organic synthesis to wastewaters treatment. Due to these great enzymes potential application, their production is being studied in submerged and solid state fermentation, using synthetic substrates and agro-industrial residues, with free and immobilized microorganisms in different supports (Songulashvili et al., 2007; Couto and Sanromán, 2005; Duran et al., 2002). As formerly mentioned several microorganisms, mainly white rot fungi, have been researched for ligninolytic enzyme acquirement, among the most studied, the genera Phanerochaete, Pleurotus, Agaricus and Lentinula can be cited (Rodriguéz et al., 1999; Wong and Yu, 1999; Espósito et al., 1991; Waldner et al., 1988; Tien and Kirk, 1983; Burdsall and Eslyn, 1974). Another interesting and low cost form to obtain ligninolytic enzymes is through solid residues extraction from edible mushrooms production (e.g.: Pleurotus ostreatus and Agaricus blazei). These fungi culture generates a great deal of solid residues rich in oxidative enzymes, mainly laccases, which can be extracted and used with different purposes. It is worth emphasized that the worldwide mushrooms production has increased, motivated by functional properties off this food type.
Figure 6. Heme peroxidase catalytic cycle.
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Peroxidases are extracellular enzymes H2O2 dependent that catalyze oxidation reaction, via 1-electron, in a great amount of compounds. Lignin peroxidases (EC 1.11.1.14) are glycoproteins with molecular mass about 40,000, presenting as prosthetic group iron protoporphyrin IX (Tuor et al., 1995). Lignin peroxidase (LiP) catalytic cycle, for example, includes the hydrogen peroxide activation and oxidized intermediates formation, as: LiP I, LiP II, LiP III, as can be seen in Figure 6. LiP (ferric state, Fe3+) is initially oxidized by H2O2 to form the compound LiP I (ferryl porphyrin cation radical, Fe4+) that is 2 electrons deficient. LiP I oxidizes the substrate, via 1-electron, forming the compound LiP II (oxi-iron intermediate, Fe4+) that can also oxidizes substrate leading the enzyme return to the native state. Hydrogen peroxide over addition leads to the formation of LiP III enzyme inactive intermediate, that can be converted quickly to the compound LiP III*, irreversible inactive enzyme form. This catalytic cycle is generally common for all heme peroxidases. As LiP, manganese peroxidase (EC 1.11.1.13) (MnP) is a heme protein glycosylated with molecular mass between 32 and 62.5 kDa (Hofrichter, 2002) that also needs H2O2 in its activation. Lignin natural oxidation mechanism involves oxidation of Mn(II) to Mn(III). Mn(III) is stabilized by chelant agents, such as oxalic acid. This chelant complex Mn(III) is highly oxidant and has a high diffusion capacity, due to its small size. Laccases (EC 1.10.3.2) are glycosylated proteins, in general dimeric or tetrameric, called blue oxidases. They have molecular mass between 60 to 390 kDa (Call and Mücke, 1997) and are called blue oxidases due to four copper atoms presence per monomer, being distributed in three redox sites (Gianfreda et al., 1999; Couto and Herrera, 2006). Copper oxidation state in native enzyme is +2 (McGuirl and Dooley, 1999). Laccases are able to oxidize a great variety of aromatic compounds hydrogen donor with concomitant oxygen and water reduction. The action spectrum of laccases can also be enlarged through the use of small molecules able to act as mediators in electron transference. These systems that use mediators, LMS (laccases mediated system), have already been applied in organic compounds degradation, in biosensors development, in biopulping, among others examples (Wesenberg et al., 2003; Bourbonnais et al., 1998; Bourbonnais et al., 1997; Trudeau et al., 1997; Collins et al., 1997). The most used mediators are presented in Figure 7. Despite of does not belong to any ligninolytic system and, indeed, be a vegetal origin enzyme, horseradish peroxidase (HRP, EC 1.11.1.7) deserves distinction among peroxidases, not only for its wide catalytic application, but also for being one of the first studied peroxidase. In 1955, Hugo Theorell won the Nobel Prize in Medicine due to his researches with redox enzymes involving HRP (Dunford, 1999). Horseradish peroxidase is the best know and better characterized vegetable peroxidase until now. HRP is also a glycoprotein with iron protoporphyrin as prosthetic radical, being its molecular mass about 40,000. This enzyme is strongly used in diagnostic kits for H2O2 determination in clinical analysis. More pure preparations are used as protein markers in cyto- and histochemistry. HRP enzyme preparations can also be used in immunological assays. The use of HRP in biosensors, in effluents remediation, in organic synthesis and therapeutic applications have already been reported (Madeira et al., 2008, Mello and Kubota, 2007; Ryan et al., 2006; Ferreira-Leitão et al., 2003; Colonna et al., 1999; van de Velde et al., 2001; Meunier and Meunier, 1985; Kedderis and Hollenberg, 1983).
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Figure 7. Chemical structure of most used mediators in LMS (Laccase mediated system). HBT (1hydroxybenzotriazole), ABTS (2,2-Azinobis(3-ethylbenzthiazoline-6-sulfonate)), Viorulic acid.
4. Biocatalyst Application in Environmental Technology 4.1. Detoxification and Valorization of Oleaginous Solid Waste The castor bean contains about 50% oil, which has special characteristics such as high viscosity, stability to heat and pressure, low freezing point, and ability to form waxy substances upon chemical treatments (Conceição et al., 2005). As consequence of this, castor bean is an oleaginous candidate for biodiesel production. In recent years, the Petrobras Research Center is developing a biodiesel production process from castor bean seeds (Khalil and Leite, 2006). After the transesterification reaction, a reject named castor bean waste is produced. This waste, extremely alkaline, presents no utility after biodiesel production and it cannot be used as animal feed. Furthermore, castor bean seeds contain a strong toxin (ricin)
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and an allergenic protein fraction (CB-1A or 2S albumin isoforms), which severely limits the usefulness of the castor meal after the oil extraction (Audi et al., 2005; Thorpe et al., 1988). Ricin toxin is a 64 kDa protein produced by castor beans (Ricinus communis). The holotoxin consists in two polypeptide chains of 32 and 34 kDa connected with a disulphide bond. The A chain (RTA) is a ribotoxin which inhibits protein synthesis in mammalian cells. The B chain (RTB) is a lectin, which binds to galactose residues on the cell surface. A single molecule of RTA in the cell cytoplasm inhibits completely the protein synthesis. The estimated lethal dose of ricin in humans is 1–10 μg/kg. (Audi et al., 2005; Rao et al., 2005). The allergenic compounds are unusually nontoxic stable proteins that exhibit an extraordinary capacity to sensitize individuals exposed to small concentrations of the dust from castor beans or the castor bean waste. The total elimination or inactivation of toxic and allergenic compounds is extremely important for the manipulation and final destination of the castor bean waste. Attempts have been made to detoxify the castor cake, byproduct of the castor oil extraction by physical and chemical methods (Anandan et al., 2005). However, biological detoxification by solid state fermentation represents an interesting alternative to the generation of biotechnological interest products at lower costs, such as enzymes. Recently, Bevilaqua et al. (2007) developed a new methodology for biological detoxification of castor bean waste and aggregate value to the residue by the production of an acidic thermo stable lipase in SSF by Penicillium simplicissimum strain, previously selected as excellent lipase producer (Gutarra et al., 2005; 2007). The wild Brazilian P. simplicissimum strain was able to grow and produce a higher lipase activity by solid state fermentation with castor bean medium compared to the production of this enzyme in babassu cake (Gutarra et al., 2005) and soy cake (Di Luccio et al., 2004). The detoxification of the castor bean medium by P. simplicissimum growth was evaluated, by Bevilaqua et al. (2007) during fermentation course up to 72h by electrophoresis SDS-PAGE (Figure 8). It was observed a gradual ricin bands reduction over time. In only 24h of fermentation, a reduction of approximately 30% in the ricin amount was verified. After 48 h the ricin bands were eliminated (Figure 8 - lanes 3 and 4). Probably, the fungus uses ricin as nutrient source, reducing it to non detectable levels and consequently shortening the waste toxicity. Allergenic potential was evaluated through mast cell degranulation. Mast cells were treated with 2S albumin fraction isolated from gel-filtration chromatography of crude extract of each sample. Mast cell degranulation, promoted during the isolation of these cells, was 30 %. Sensitized cells treated (positive control) and non-treated (negative control) with 2S albumin pool showed 64.6 and 33.8 % of mast cell degranulation, respectively. The 2S albumin fraction isolated from natural sample showed 57 % mast cell degranulation while after 72h of fermentation the samples presented fewer cell degranulation. Besides ricin elimination of the castor bean waste, the fermentation by the fungus P. simplicissimum showed almost 16 % allergenic potential reduction.
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Figure 8. SDS-PAGE of proteic extracts of castor bean waste. Lane M- protein MW marker; lane 1- nonfermented waste; lane 2- 24h fermented waste; lane 3- 48h fermented waste; lane 4- 72h fermented waste. (Bevilaqua et al., 2007)
The castor bean waste constitutes an interesting alternative for lipase production, considering a higher production achieved, compared with the other cited residues, being able to increase the productivity after the optimization process. Moreover, castor bean waste is a reject that, in consequence of its high rate of toxicity and allergenicity, normally is not used for other finalities, as babassu and soy cake, which can be used as animal feed, for example. Therefore, it was possible to aggregate value to a high undesirable waste, by the production of an enzyme with greater biotechnological potential. So, it was developed a new low cost methodology for biocatalyst production of high aggregate value and, simultaneously, the detoxification of an agro industrial waste.
4.2. Detoxification of Poultry Slaughterhouse Solid Waste Voluminous poultry feather amount are generated by poultry industry, since about 5-7% of poultry weight is constituted by feathers. An industrial poultry slaughterhouse can reach a production of 50,000 poultry, what corresponds to the generation of nearly 2 or 3 tons of feathers (Dalev, 1994; Onifade et al., 1998). Considering that about 90% of the feather weight is compound by keratin and still considering the high consistency and, consequently, the slow degradation of this material in natural environments, feathers become pollutant, where sulphurous compounds production
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with an extremely nasty smell generates environmental problems (Onifade et al., 1998; Sangali and Brandelli, 2000). The incineration process in suitable installation, as palliative solution to banish these biological residues is economically inefficient (Dalev, 1994). Due to feathers being a natural source with high protein content (thereabout 750g of feather / kg of crude protein), they have been profited for feather meal production through treatments, mainly chemical and thermic ones, for after incorporation as feed supplement in animals feed (Dalev, 1994). This process results in some essential aminoacids, such as methionine, lysine, hystidine and tryptophan (Williams et al., 1990; Dalev et al., 1997). However, employs expensive equipments for long periods (8-12 h), besides spent intensive energy. The difficulties here reported, have motivate the search for development of non-pollutant process (Onifade et al., 1998) for feather meal production. Elmayergi and Smith (1971) showed that is possible the modification in feather keratin structure by microorganisms cultivation, as Streptomyces fradiae, and by keratinolytic enzymes therein produced which can facilitate the animal feed digestibility. Besides feather meal, the microbial biomass can also be used as complement or additive in protein enrichment of animal feed. Higher amounts of lysine, methionine and arginine were detected in fermented feathers when compared to the ones found in non fermented feathers concluding that not only feather keratin but also microbial biomass can be employed as protein sources (Onifade et al., 1998). Despite the limited available information about feathers biodegradation by microorganisms, this biological treatment represents an alternative for the increase in feather utilization as protein supplement in animal feed and for aminoacid production in its pure form (Williams et al., 1990). Besides these benefits, a significant increase in fowl growth was observed when feather lysed, belonging to bacterial fermentation were added to these animals feed, instead of commercial or non treated feathers (Lin et al., 1992). Lin et al. (1992) isolated a strain of Bacillus licheniformis PWD-1 able to grow using feathers as primary organic substrate for carbon, sulphur and energy supply. The feathers biodegradation by this bacterium represents an improvement in feather employment as protein source, for example. Searches carried out with keratinases produced by Aspergillus fumigatus indicate that these enzymes are able to degrade keratin in natura and they have potential to be used in biotechnological process in feathers degradation (Santos et al., 1996). Azeredo et al. (2001; 2006) observed Streptomyces sp. 594 growth using different keratin substrates, as sheep wool, chicken feather and feather meal indicating the ability of this microorganism binding itself to these substrates and degrade them through keratinolytic proteases. The degradation capacity of feather structures was investigated on solid and in liquid media, after 15 and 7 days of incubation at 30 ºC, respectively. Casitone-molasses and mineral media, both containing whole poultry feather, were used and an autoclaved and noninoculated poultry feather was applied as control. The feather degradation from submerged cultivation was microscopically observed using Nomarski differential interference contrast (DIC) technique. Comparing to the autoclaved and non-inoculated feathers (control) (Figure 9a, 9c), feather barbules and rachis were hydrolyzed by Streptomyces sp. 594 in basal agar medium and casitone-molasses medium (Figure 9b, 9d).
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a
c
b
d
Figure 9. Poultry feather degradation by Streptomyces sp. 594. In mineral agar medium (a-control; binoculated) and in complex liquid medium (c-control; d-inoculated), 40X magnification. (Azeredo et al., 2006).
Relative results to keratin hydrolysis capacity were also observed with the Chryseobacterium sp. kr6 strain in mineral medium added with feather. Feather barbules were completely degraded and rachises were also attacked by the bacteria. The isolate was able to disintegrate thoroughly the upright feathers (Riffel et al., 2003). The content of keratin total degradation by microorganisms only occurs after its denaturation by disulphide bond cleavage that represents the main extraordinary stability source and proteolytic digestion resistance through sulphitolysis. The phenol group formation
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by Vibrio kr2 strain suggests an enzyme presence, the disulphide reductase, in the cultivation medium added with feather keratin (Sangali and Brandelli, 2000). The first and only report in literature about two enzymes co-operative that results in efficient keratin degradation was discussed (Yamamura et al., 2002). A Stenotrophomonas sp. D1 strain, isolated from a deer horn, produced two types of enzymes. These two enzymes action working co-operatively resulted in a greater keratinolytic activity. The biochemical properties results suggest that this protease is a serine protease and by the reduced protein of disulphide bonds it can be a type of disulphide reductase (Yamamura et al., 2002). Due to the recent tendency in clean technology development, the proteases, especially keratinases, are potential candidates for environmental applications like in feather degradation.
4.3. Coffee Residues Detoxification Coffee is a very important agro-industrial commodity in the world and its production generates high amounts of by-products. It is estimated that only approximately 6% of the fresh fruit is used in a preparation of coffee as a drink, and the remaining 94% is water and by-products of the production process (Pandey et al., 2000b; Soccol and Vandenbergue, 2003). The powder coffee can be produced by dry or wet process and generates the husk or pulp solid wastes, respectively. These two residues are rich in nutrients showing high quantities of carbohydrates, proteins and fats. They also show in its composition antinutritional factors such as caffeine, polyphenols and tannins (Pandey et al., 2000b). As these compounds cause low feed intake, poor protein digestibility and nitrogen retention, the use of coffee wastes as animal food is restricted. These compounds are also anti-physiological factors restricting microbial growth and thus the solid coffee residues are not well explored constituting a considerable source of environmental pollution. Detoxification methodologies had been tested to eliminate or reduce anti-nutritional factors presented in coffee wastes. Physical and chemical methods were successful but were very expensive and non economical feasibly. In this way, biological methodologies had received additional attention such as strategies based on microorganisms’ growth by solid state fermentation. In this strategy, the microorganism grows in solid wastes and removes or at least degrades to a very low level the toxic compounds by the action of different enzymes as tannases and caffeinases. Strains of filament fungi, mostly from the genera Penicillium and Aspergillus, were isolated from coffee samples and its by-products. These strains when growing in liquid medium showed the ability to degrade caffeine (Roussos et al., 1995). The biological detoxification of coffee pulp and husk by solid state fermentation using filamentous fungi with simultaneous nutrient enrichment had been a subject of several scientific works (Krishna, 2005; Pandey et al., 2000b). The growth of phytase producing microorganisms could enhance the digestibility of these residues (Pandey et al., 2001). Brand et al. (2000) studied the detoxification of coffee husk in solid state fermentation using fungal strains. A total of 14 fungal strains, were screened from their ability to grow on coffee husk extract agar (CHA), a medium prepared by cooking coffee husk in distilled water. All strains were able to grow on coffee husk extract agar, showing resistance to toxic factors therein
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presented. The fungi Rhizopus arrhizus LPB-79, Phanerochaete chrysosporium BK and Aspergillus sp. were the strains which showed high quantities of biomass in coffee husk extract agar medium and were used for coffee husk detoxification in solid state fermentation. The Aspergillus sp. showed best degradation results, reaching removal of 92% of caffeine and 65 % of tannins. This result was already expected as the Aspergillus sp. strain tested was the unique strain isolated from coffee husk and the one that showed best growth on coffee husk extract agar medium. The upgrading of nutritional quality of coffee husk was studied through the growth of Aspergillus niger LPBx strain, isolated from coffee husk in Brazil, by solid state fermentation of this residue (Brand et al., 2001). Fractional and factorial designs were carried out to optimize process parameters as moisture, aeration rate and inoculum size. Results showed that moisture and aeration rate were significant factors for the degradation of caffeine and tannin. Best results were achieved with 30mL/min aeration and 55% of initial moisture, reaching degradation of 90% of caffeine and 57 % of tannins. After fermentation, a 2-time increase in protein contents of the coffee husk was achieved. The detoxification and proteinenrichment of coffee husk make possible the utilization of this residue as animal feed. Brand et al. (2002), using the same strain, showed the relationship between caffeine degradation and fungus respiration. The kinetic study showed that caffeine degradation was related to fungus development, to protein synthesis and also with the consumption of reducing sugars present in coffee husk. Thus, the authors suggested that the reduction of caffeine could be related to its consumption as nitrogen source by the fungus. Detoxification of toxic compounds from coffee wastes was also reached using bacteria as biological agents. Ulloa Rojas et al. (2003) used three different biological treatments: ensiling adding sugar cane molasses, aerobic decomposition and aerobic bacterial inoculation with cellulolytic bacteria. Ensiling for 2 and 3 months of coffee pulp reduces cellulose, polyphenols and tannins but caffeine contents remain the same. With this treatment protein and fats levels tend to increase. Aerobic decomposition up to 42 days also increases protein and fat levels and reduces cellulose, phenols, caffeine and tannins contents by increasing treatment time. Caffeine was totally degraded after 14 days. Aerobic bacterial inoculation with Bacillus sp. strains increased the protein content of coffee pulp after 21 days and cellulose, total phenols, tannins and caffeine contents reduced with time of bacterial degradation. Both the aerobic decomposition and the aerobic bacterial degradation appeared more suitable to improve the nutritional quality of coffee pulp than the ensiling. These results showed that biological treatment envolving biocatalystis is an efficient methodology to reduce toxic compounds of coffee waste as tannins, caffeine and polyphenols and to increase mostly protein contents. These composition changes in coffee wastes add nutritional values to this residue enhancing its applicability as animal feed. Strain that shows ability to degrade phenolic compounds in coffee wastes could also be utilized for the removal of these compounds when presented in others agro-industrial wastes as olive oil wastes (cake, olive mill and semi-solid residues) (Saavedra et al., 2006; Linares et al., 2003; D’Annibale et al., 2006).
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4.4. Cassava Residues Detoxification Cassava (Manihot esculenta Crantz) is widely consumed as human food mostly in tropical countries in Asia, Africa and Latin America and is largely produced in about 88 countries (Soccol and Vandenberghe, 2003; Pandey et al., 2000c). High production and consumption are associated with cassava remarkable capacity to adapt to various environmental conditions: even in poor soils under dry conditions it still produces edible roots. Cassava is also a valuable low-cost source of calories. Despite of these advantages, cassava shows low levels of proteins and essential minerals and vitamins in its composition. Some varieties of cassava shows high concentrations of cyanogenic glucosides (linamarin and a small amount of lotaustralin), which can be hydrolyzed to hydrocyanine acid by endogenous linamarase released from the plant tissue during the roots processing. Cassava roots also contain tannic acid and this compound can give dark color to processed products and affect protein digestibility (Oboh and Akindahunsi, 2003). Depending on cassava variety, cultivation and environmental conditions, different quantities of cyanide could be achieved in cassava parenchyma and toxicity must be reduced to safe levels before consumption (Oluwole et al., 2007; Cardoso et al., 2005). Its known that water stress enhances cyanide contains. The World Health Organization (WHO) has set the safe level of cyanogens in cassava flour at 10 ppm (FAO/WHO, 1991). Cassava root is industrially processed to obtain flour, starch and natural fermented food as gari and fufu (Agbor-Egbe and Lape Mbome, 2006). They can be produced by different methods which vary depending mainly on the region and the choice is based in economical conditions. Methods which use crushing (more expensive) are very effective in removing cyanide due to the intimate contact in the finely-divided wet parenchyma between linamarin and linamarase, which promotes rapid breakdown of linamarin to hydrogen cyanide gas that escapes into the air. Cheaper methods, commonly used in eastern and southern Africa, as sun drying or heap fermentation showed low cyanide removal (approximately 25–33% retention of cyanide). In these methods peeled roots are usually cut in half longitudinally and most plant cells remain intact, with the linamarin stored inside the cell separately from the linamarase, located in the cell walls. Heap fermentation is better than sun drying mainly due to a solid state fermentation process with endogenous microorganisms that reduces cyanide contents. Combined methods for roots processing cause satisfactory removal of cyanogens. Flour produced by crushing and sun drying showed retention of 1.5–3.2% in cyanide contents. The efficiently removal of cyanides depends on the method and mainly on cyanide concentration on root because even better methods as the ones which produce flour in Brazil and gari in West Africa can fail if the level of cyanide in the root is higher than 222 ppm (Cardoso et al., 2005; Agbor-Egbe and Lape Mbome, 2006). Based in these researches, attempts have been done to improve products as gari and flour employing solid state fermentation with pure strains of filamentous fungi or Saccharomyces cerevisiae during manufacture process. This procedure can reduce cyanide contents and also enhance proteins levels. Oboh and Akindahunsi, (2003) showed approximately a 2-time increase in protein contents and a 2-time decrease in cyanide levels of gari and flour using pure strain of Saccharomyces cerevisiae. Solid state fermentation of the cassava pulp to produce gari and flour with Aspergillus niger also showed an increase of protein contents and
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a reduction of cyanide levels when compared with non-fermented or natural fermented products (conventional method to produce gari and flour) (Oboh et al., 2002). From cassava processement high quantities of waste is produced. They can be separated in liquid or solid wastes and by type and its proportion depends on the process method. Solid residues can be separated mainly in peel, fibrous residue (pulp waste or bagasses), and carbohydrate rich pressing slurry (Ubalua, 2007). The peel constitutes 20 - 35% of the total weight of the tuber and shows high quantities of cyanides when the bagasses show no cyanide contents (Pandey et al., 2000c; Ugwuanyi et al., 2007). Total solid cassava waste shows high quantities of starch in its composition but has poor protein contents. High cyanide contents and low proteins values make this abundant residue unattractive for animal nutrition use. Animal consumption of non- or poor detoxified cassava has been associated with chronic toxicity, resulting in lowered egg production in poultry and poor growth of livestock. Solid cassava waste is generally discarded in environment as landfill without any treatment. Natural degradation occurs slowly and causes strong and bad smell. The large amount of cassava waste which has no interesting applicability represents serious environmental problems causing cyanide contamination of water bodies and environment (Soccol and Vandenberghe, 2003). So, fermentation has been identified as one of the less expensive detoxification means and protein quality increase of cassava waste (Ubalua, 2007). Ensiling has been employed to reduce cyanide contents and to promote nutritionalenrichment. During ensiling, anaerobic bacteria multiply rapidly, predominating strains of Lactobacillus that produce large amounts of lactic acid, reducing the residue pH at the end of the process to 4.0. Other species also grow during ensiling and many products can be formed. Ensiling of cassava waste could diminish the cyanide level allowing the product be used subsequently as animal feed (Ubalua, 2007). Oboh (2006) studied the removal of cyanides from cassava peels using solid state fermentation. Cassava peels were fermented by adding cassava wastewater (liquid squeezed out of the fermented parenchyma mash) as inoculum since this liquid residue shows high quantities of microorganisms able to hydrolyze cyanides in its composition. Two types of treatment were tested: (a) wastewater obtained by the natural fermentation of cassava pulp; and (b) wastewater obtained by fermentation with pure strains of Saccharomyces cerevisiae and two bacteria, Lactobacillus delbruckii and Lactobacillus coryneformis. Nutritional value and cyanide contents were analyzed in non-, natural and inoculated fermented cassava peels. Analysis of the dried fermented peels showed a 2.6 -time increase in the protein content of inoculated fermented cassava peels when compared with unfermented ones. Furthermore, this treatment showed a 7.2 and 1.3 -time decrease in cyanide and phytate (probably by the production of different enzymes as cyanidases and phytases) content, respectively when compared with the unfermented cassava peels. Natural fermented cassava peel also promotes protein enrichment and reduction in antinutritional contents, but the inoculated process which uses wastewater obtained by cassava pulp fermentation with S. cerevisiae, Lactobacillus delbruckii and L. coryneformis showed better results. Fermentation of cassava peel inoculated with cassava wastewater could be a good protein source in livestock feeds. The detoxification of cassava waste was achieved by the growth of linamarase producers microorganisms. Ugwuanyi et al. (2007) isolated strains of Bacillus sp. in waste treatment process using thermophilic aerobic digestion which showed linamarase activity.
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4.5. Wastewater Treatment with High Oil and Grease Levels The steadily rising demand for food products has required an increase in productivity from food industries, and thus an increase in the numbers of dairies, slaughterhouses and meat processing plants in many countries. Among the food industries, the dairy industry is the most polluting in volume (generating from 0.2 to 10 m3 of effluent per m3 of processed milk) in regards to its large water consumption (Vourch et al., 2008). In slaughterhouses, the consumption of water per slaughtered animal varies according to the animal and the industryspecific process employed, ranging from 1.0 to 8.3 m3. Most of this is discarded as wastewater, with 0.4 to 3.1 m3 of water per slaughtered animal being reported in the literature (Caixeta et al., 2002). Wastewaters from food processing industries such as those coming from dairies (Danalewich et al., 1998; Cammarota et al., 2001; Jung et al., 2002; Marshall and Harper, 1984; Omil et al., 2003), slaughterhouses (Batstone et al., 1997; Martínez et al., 1995; Massé et al., 2001, 2003; Ruiz et al. 1997; Caixeta et al., 2002; Fuchs et al., 2003; Wang and Banks, 2003; Torkian et al., 2003) and fat refineries (Faisal and Unno, 2001; Saatci et al., 2003) are rich in biodegradable organic molecules and nutrients and usually contain high levels of fats and proteins that have a low biodegradability coefficient. If not treated, they cause gross pollution of land and water with their high biochemical oxygen demand (BOD) and chemical oxygen demand (COD). The technology of lipid biodegradation has not yet been fully exploited in the processing of tons of one of the major organic material present in wastewater streams; hence the restrictions on its disposal. In countries where food habits result in a large amount of residual fat and oil in wastewater, it has become increasingly difficult to fulfill the discard requirements (Tano-Debrah et al., 1999). Conventional methods for wastewater treatment include aerobic processes such as aerated lagoons, activated sludge (Carta-Escobar et al., 2005; Shack and Shandhu, 1989; Stephenson, 1989), membrane sequencing batch reactors (Bae et al., 2003), trickling filters (Walsh et al., 1994) and rotating biological contactors (Radick, 1992). However, the energy requirements for the aeration in these installations are high and problems such as bulking and excessive biomass growth frequently occur under these conditions (Timmermans et al., 1993). In the last two decades anaerobic reactors have been increasingly applied for treatment of these wastewaters (Omil et al., 2003) by means of upflow anaerobic sludge blanket (UASB) reactors (Hansen and Hwang, 1990; Hawkes et al., 1995; Rico et al., 1991; Martínez et al., 1995; Kim et al., 2004), hybrid UASB reactors (Ozturk et al., 1993), expanded granular sludge bed (EGSB) reactors (Petruy and Lettinga, 1997), as well as others based on anaerobic filters (Méndez et al., 1989; Veiga et al., 1994; Viraraghavan and Kikkeri, 1990), have been reported. These studies demonstrated that aerobic and anaerobic treatments can be used effectively for these effluents. However, it is necessary to reduce the concentration of fat, oils and proteins or to eliminate these materials altogether, in order to enable the biological treatment to proceed without any inhibition of the biological reduction of organic matter in wastewater. A large number of pretreatment systems (grease-trap, tilted plate separators, dissolved air flotation systems and physical-chemical treatment) are employed to remove oil and grease (O&G) from these wastewaters prior to the main treatment process itself, which is generally
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of biological nature. However, the cost of such reagents is high, the removal efficiency of dissolved and/or emulsified O&G is low and extremely problematic sludge is produced (Kárpáti et al., 1995; Tano-Debrah et al., 1999; Willey, 2001). The O&G not retained in the pretreatment systems enter into the biological treatment system, becoming a considerable nuisance, especially in conventional mesophilic processes (Massé et al., 2001). These O&G are associated with several problems, including the reduction of the cell-aqueous phase transfer rates (substrates, products and oxygen) through the formation of a lipid coat around the biological floc (Chao and Yang, 1981; Saxena et al., 1986; Grulois et al., 1993; Lemmer and Baumann, 1988; Rinzema et al., 1994). In addition, filamentous microorganism blooms (bulking) and floating sludge with undesirable physical characteristics may develop. Furthermore, poor activity associated with excessive amounts of O&G in the wastewater hinders sedimentation and causes losses of biomass through the reactor’s outflow (Bae et al., 2003; Danalewich et al., 1998; Jenkins et al., 1993). These detrimental effects are further associated with clogging and the emergence of unpleasant odors, and are frequently associated with a reduction in the efficiency of treatment stations (Carta-Escobar et al., 2005; Grulois et al., 1993; Rinzema et al., 1993; Perle et al., 1995; Vidal et al., 2000; Martínez et al., 1995). Sayed et al. (1988), studying the anaerobic degradation of various fractions of slaughterhouse wastewater, verified that the process-controlling factor was the liquefaction of colloidals adsorbed on the bacteria and the hydrolysis of coarse suspended solids entrapped within the biomass bed. Petruy and Lettinga (1997) showed that 70% of lipids were adsorbed by the granular sludge, within approximately one day, and thereafter, the remaining lipids were slowly converted into methane gas in an anaerobic reactor. Jeganathan et al. (2006), using upflow anaerobic sludge blanket (UASB) reactors to treat a complex oily wastewater from a food industry, reported that although approximately 75% of COD was degraded (which corresponded to the observed biogas yield) for an organic loading rate of about 2.5 kg COD/m3.day, system performance deteriorated sharply at higher loading rates. An increase in loading to 5 kg COD/m3.day caused O&G accumulation in sludge and increase in scum production which reduced degradation to 40–50%. These authors also reported that accumulation of O&G in the biomass was the critical parameter governing the high-rate anaerobic reactor performance and further suggest the need for periodic reseeding of anaerobic reactor systems treating oily wastes, since the loss of sludge in the bed, due to washout, increased the O&G loading to the biomass and failure occurrence. The biodegradation of lipids is difficult due to their low bioavailability (Petruy and Lettinga, 1997). Fats in wastewaters produce glycerol and LCFA (saturated fatty acids with 12 to 14 carbon atoms and unsaturated fatty acids with 18 carbon atoms) during the hydrolytic step. Glycerol was found to be a non-inhibitory compound (Perle et al., 1995). However, LCFA have been reported to inhibit the activity of various microorganisms (Angelidaki and Ahring, 1992; Hanaki et al., 1981; Koster, 1987; Rinzema et al., 1994). They also decrease the availability of adenosine triphosphate (ATP) (Hanaki et al., 1981; Perle et al., 1995). The inhibitory effect increases with the number of double bonds and cis-isomers that are abundant in natural lipids (Rinzema, 1988). As in the case of volatile fatty acids (VFA), the toxicity of LCFA seems to be related to the unionized form of these acids, namely the long chain free fatty acids (LCFFA). In upflow anaerobic sludge blanket (UASB) or the
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expanded granular sludge bed (EGSB) reactors, is the adsorption of LCFA that induces sludge disintegration, flotation and washout (Cavaleiro et al., 2007). Also transport limitation phenomena, caused by LCFA accumulation onto the sludge, was found to have an important contribution to the observed lag phases preceding methane production, generally reported to be ascribed to mechanisms of cell wall damage or to long term acclimation (Pereira et al., 2005). It was found that the observed temporary decrease in the methanogenic activity after the contact with LCFA is a reversible phenomenon, being eliminated after the conversion to methane of the biomass-associated LCFA, suggesting that sequencing accumulation/ degradation steps could promote a sustainable biogas production from LCFA and the anaerobic treatment of a fat rich dairy wastewater is enhanced when repeated pulse feeding is applied (Cavaleiro et al., 2007). The biological treatment of fats and oils under thermophilic conditions is advantageous due to the favorable changes in most physical properties of hydrophobic compounds at high temperatures. These substances may become more accessible to microorganisms and their lipolytic enzymes when the process occurs under thermophilic conditions (Becker et al., 1999; Thomas, 1987). Despite the increased efficiency and decreased residence times that can be achieved under thermophilic conditions (in comparison with mesophilic processes), largescale application of this technology poses serious challenges for treatment systems. Namely, expensive modifications involving the inclusion of heating systems, maintenance and temperature control are frequently required. The application of a pretreatment process to hydrolyze and dissolve fats may improve the biological degradation of fatty wastewaters, accelerating the process and reducing time. Consequently, the treatment of effluents with high fat contents from several origins is a new and promising actuation area for lipases. Candida rugosa lipases were used in the treatment of domestic wastewaters and in the cleaning of sewer systems, cesspools and sinkholes (Jaeger and Reetz, 1998). The use of lipases from Pseudomonas aeruginosa in effluents of restaurants (Dharmsthiti and Kuhasuntisuk, 1998) has also been investigated. Lipases have also been used to accelerate the biodegradation of polymers (Marten et al., 2003; Sivalingam et al., 2003; Takamoto et al., 2001) and of slurries from oil-well perforations containing synthetic esters emulsified in water (Aliphat et al., 1998). Lipases can also be used for oil degradation on soil, deriving from spilling, or conversion of polyesters residues into useful products, such as non-esterificated fatty acids and lactones (Piras et al., 1994). In addition, the fatty effluents treatment, as the ones from slaughterhouse and dairy industries, also represents a wide field for lipase application. There are research reports and patents that have described the use of microorganisms and/or enzymes pools developed in laboratory for the biological treatment of effluents with high fat and oil concentrations. For example, lipases from Candida rugosa for fat removal in equipment of effluent treatment plants and other bioorganic catalytic formulations that rapidly break down organic contaminants including fats, oils, and greases. However, before these all products can be widely applied, it needs to be examined whether there are hazardous species present and whether they have any adverse environmental effects, especially when used in places with conditions that differ considerably from the conditions under which the microorganisms were isolated.
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Wakelin and Forster (1997) investigated the microbial treatment of waste from fast-food restaurants for the removal of fats, oils and greases. They cultivated pure and mixed microbial flora (known to produce lipases and other enzymes), and found that Acinetobacter sp. was the most effective of the pure cultures, typically degrading 60 to 65% of the fatty material, which initial concentration had been 8 g/L. The strain P. aeruginosa LP602 was tested for its ability to treat this type of wastewater (Dharmsthiti and Kuhasuntisuk, 1998). When these cells were added to the wastewater, the BOD was reduced by 94% after an 8-day incubation period. The lipid content in the wastewater was also rapidly reduced, and all the lipid content was removed within the first 5 days of treatment. Crude lipase from P. aeruginosa LP602 (3.5 U/mL) was added to this same lipid-rich restaurant wastewater in a ratio of 1:1 and incubated with shaking at 37°C. The lipid content (approximately 200 mg/L) was reduced by 70%, to less than 10 mg/L, during the first 24 h and was not detected after 48 h. Tano-Debrah et al. (1999) developed an inoculum with high fat and oil degrading activities that consisted in a mixed culture of 15 bacterial isolates from various fatty wastewater samples taken from grease-traps of restaurants in Japan. All isolates were able to degrade various fats and oils to some degree. However, the extent of degradation varied for each fat/oil tested, being lower for shea fat (fat obtained from nuts of the tree Butyrospermum parkii). The variation in the composition of the fat/oil mixture caused variations in the development of the inoculum bacteria. Massé et al. (2001) evaluated the effect of alkaline and enzymatic pretreatments on the solubilization and size reduction of fat particles in slaughterhouse wastewater. The average particle size was reduced to 73% of the initial average particle size at NaOH concentrations ranging from 6 to 12 g/L. The high doses of NaOH required and the resulting increase in pH rendered alkaline pretreatment a poor choice for a hydrolysis pretreatment of fat particles. However, the pretreatment with pancreatic lipase PL-250 reduced the average particle size to a maximum of 60% from initial particle size. The bacterial lipase LG-1000 was also efficient in reducing the average fat particle size, but high doses (greater than 1000 mg/L) were required to obtain a substantial reduction after 4 h of pretreatment. No particle size reduction or changes in soluble COD were noted after 4 h of pretreatment with the plant lipase EcoSystem Plus. It was thus concluded that pancreatic lipase PL-250 was the best known pretreatment for hydrolyzing fat particles in slaughterhouse wastewater. Mongkolthanaruk and Dharmsthiti (2002) evaluated a mixed culture composed of P. aeruginosa LP602, Acinetobacter calcoaceticus LP009 (both lipase-producing bacteria) and Bacillus sp. B304 (an amylase and protease producing bacterium) to lower the biochemical oxygen demand (BOD) value and lipid content of lipid-rich wastewater. The BOD and the lipid content were reduced from 3600 mg/L and 21,000 mg/L, respectively, to less than 20 mg/L within 12 days under aerobic conditions. Enzymes and pure cultures have also been used to increase hydrolysis during or prior to biological treatment process (Aoki and Kawase, 1991; Cail et al., 1986; De Felice et al., 2004; Lagerkwist and Chen, 1993; Lanciotti et al., 2005; Rintala and Ahring, 1994). Such pre- or co-treatments methods generally consist in the cultivation of lipase-producing microbial strains in the effluents. Cail et al. (1986) tested an enzymatic mixture containing protease, amylase, cellulose, lipase and Bacillus subtilis spores on wool scouring wastewater
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with high lipid content; this mixture increased the COD reduction from 59% in the control to 78%, increased grease removal from 47% to over 70%, and improved solids reduction from 34% to over 70%. Rintala and Ahring (1994) however reported that the addition of an enzymatic supplement (lipase, cellulase and protease) into an anaerobic reactor treating household solid waste did not improve reactor performance relative to the control (without enzymes). De Felice et al. (2004) cultivated the yeast Yarrowia lipolytica ATCC 20255 on wastewater from an olive oil mill under batch culture conditions. They found that the yeast was capable of reducing the COD value (100 to 200 g/L) by 80% in 24 h and producing a useful biomass of 22.45 g/L as single cell protein and lipases. Similar results were described by Scioli and Vollaro (1997), who reported lipolytic activity of 770 U/L in the fermentation medium after 24 h. Lanciotti et al. (2005) evaluated the ability of several Yarrowia lipolytica strains to grow in olive mill wastewater and to metabolize its lipid fraction. All strains studied were able to produce extracellular lipases. Some strains, in addition to a high lipase activity, induced a COD reduction with respect to the uninoculated wastewater (control) ranging from 20 to 40% after 72 h at 25oC. The effect of bioaugmentation with an anaerobic lipolytic bacterial strain (Clostridium lundense DSM 17049T) on the anaerobic digestion of restaurant lipid-rich waste was studied by Cirne et al. (2006) in batch experiments with a model waste containing 10% lipids (triolein). A higher methane production rate was observed in the experiment with the presence of the bioaugmenting lipolytic strain under methanogenic conditions. The levels of palmitate and stearate were also higher until day 15, indicating that the bioaugmentation strategy improved the hydrolysis of the lipid fraction. Jordan and Mullen (2007) investigated the addition of commercial extracellular enzymes (lipase, pectinase, cellulase, alcalase e laccase) to dairy wastewater sludge (DWS) in an attempt to accelerate the degradation of the associated organic matter. Preliminary results indicated that the organic material was stabilized within 9 h and that the used enzymes would, therefore, improve the efficiency of a waste management plant, if such a system was employed. The effectiveness of a commercial inoculum for degrading a dairy wastewater with high fat content was evaluated by Loperena et al. (2007) and compared with an activated sludge inoculum from a dairy wastewater treatment pond. Both inocula reached similar chemical oxygen demand removal in batch experiments (78% and 81% for commercial and activatedsludge inocula respectively). The activated-sludge inoculum produced more CO2 (1.1 and 1.8 g CO2 produced/g COD removed) than the commercial inoculum suggesting that the activated-sludge inoculum allowed a greater degree of mineralization of the effluent. The higher population diversity observed in batch reactors inoculated with activated-sludge, indicated that microorganisms from this inoculum were well adapted and may have had synergic activity for the degradation of the dairy effluent. When the bioreactor was operated with the commercial inoculum in continuous mode, according to its microbial growth kinetics, other microorganisms became predominant. These results showed that inoculated microorganisms did not persist in the open system and periodic addition of microorganisms may be needed to achieve a high performance treatment.
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The impact of the enzymatic hydrolysis of fat particles on the efficiency of a downstream anaerobic digestion process was evaluated by Massé et al. (2003). Slaughterhouse wastewater containing fat particles was pretreated with 250 mg/L of pancreatic lipase PL-250 and delivered to an anaerobic sequencing batch reactor operated at 25oC. Approximately 35% of the neutral fat was hydrolyzed during pretreatment. However, the pretreatment presented only a small overall effect on the fat particle digestion, marked by a decrease of about 5% (3 h) in the digestion time to achieve 80% of reduction in the neutral fat and LCFA concentrations. Several works in Biotechnology Center of Federal University of Rio de Janeiro, indicate the use of enzymatic preparations from solid state fermentation as an excellent alternative for the treatment of fatty effluents derived from food industry. (Cammarota et al., 2000; 2001; 2003; Jung et al., 2002, Leal et al., 2002; 2006; Cammarota e Freire, 2006, Rosa et al., 2006; Valladão et al., 2007, Damasceno et al., 2008). Leal et al. (2002) used an enzymatic extract, produced by the Penicillium restrictum fungus strain under SSF conditions, with a high level of lipase activity for the treatment of dairy wastewater with high fat contents. The wastewater was treated with 10% (v/v) enzymatic extract of 2.1 U/mL of lipase activity for 12 h at 35oC, without agitation. Both raw and hydrolyzed effluents were subsequently submitted to an anaerobic biological treatment at 35oC in bench scale. The COD removal efficiency for hydrolyzed effluents (80 to 95%) was higher than those obtained for raw effluents (without enzymatic pre-hydrolysis; 19 to 55%) at different O&G concentrations, indicate that the integrated process may be applied to treat wastewater with fat content as high as 1200 mg/L through anaerobic processes with no operating problems and with high COD removal efficiency and relatively low treatment times, which contribute to a reduction in the reactor volume. Cammarota et al. (2001) evaluated the efficacy of an enzymatic hydrolysis stage in dairy industry wastewater prior to the anaerobic biological treatment. The authors employed a solid enzymatic preparation with high Penicillium restrictum lipase activity (21 U/g). The process was tested in an upflow anaerobic sludge blanket reactor (UASB) operated for a period of 182 days at 33oC with an average organic load of 4.0 kg COD/m3.d. During this period, the O&G content of the wastewater was gradually raised (from 203 to 868 mg/L) until it reached the reactor limit in the conditions under study. The reactor limit was defined as the point when operational problems (formation of scum, biomass flotation and loss of COD removal efficiency) became so immense that had the reactor continued running under the same conditions, it would have collapsed. The evolution of total COD removal efficiency and the O&G affluent and effluent concentration data throughout the operational period from this study are presented in Figure 10. The results provide a clear demonstration of the effects of O&G accumulation in the reactor. Initially, the reactor operated with low loads of O&G (0.25 kg O&G /m3•d or 203 mg O&G/L on average) in the feeding mixture and presented total COD and O&G removal efficiencies of 91 and 82%, respectively. However, the removal efficiencies diminished drastically when the reactor received wastewater containing high levels of O&G (868 mg/L on average), reaching values on the order of 50% for COD and 40% for O&G on the 182nd day (data before vertical line in Figure 10). When wastewater pre-treated with 0.1% (w/v) of solid enzymatic preparation began to be included in the feed mixture, the reactor’s performance improved rapidly; within approximately 15 days, the
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average COD and O&G removal efficiencies had risen to their initial values of 92 and 89%, respectively (data after vertical line in Figure 10). Leal et al. (2006) also evaluated the effect of an enzymatic pre-hydrolysis stage (0.1% (w/v) of a solid enzymatic preparation (SEP) containing Penicillium restrictum lipase) on the anaerobic treatment of dairy wastewaters with different fat contents (200, 600 and 1000 mg of O&G/L) in bench UASB reactor. The UASB reactor fed with the enzymatically pre-treated effluent was able to support increments in the O&G contents in the effluent up to 1000 mg/L. When the reactor was fed with crude effluent with O&G concentration at 600 mg/L, problems manifested, such as loss of efficiency, biomass washout and fat accumulation in the sludge. ( A)
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Time of operation (days) Figure 10. Evolution of total COD removal efficiency (A) and oil and grease affluent and effluent concentrations (B) observed while monitoring the upflow anaerobic sludge blanket (UASB) reactor treating crude dairy wastewater (data before vertical line) and dairy wastewater pre-hydrolyzed enzymatically (data after vertical line), with a hydraulic retention time of 20 h at 33oC. (From Cammarota et al., 2001)
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Jung et al. (2002) evaluated the treatment of dairy wastewater containing different O&G contents in batch activated sludge systems with and without (control) an enzymatic prehydrolysis stage, which entailed a 0.2% (w/v) of fermented babassu cake containing P. restrictum lipases (11 U/g) for 8 h at 30oC. When the O&G concentration in the control bioreactor was increased (400, 600 and 800 mg/L), the COD removal efficiency decreased (86, 75 and 0%, respectively). However, in the reactor fed with pre-hydrolyzed wastewater, the COD removal efficiency was maintained (93, 92 and 82%, respectively). The initial and final COD levels for the bioreactors treating crude (A) versus pre-hydrolyzed wastewater (B) are presented in Figure 11. The bioreactor treating the pre-hydrolyzed wastewater maintained COD removal efficiencies within the range from 82 to 93% (Figure 11 B). However, at increased O&G concentrations, the COD removal capacity of the control bioreactor was challenged, such that the final COD levels in the last stage of the experiment (800 mg O&G/L), at approximately 170 days, were higher than the initial COD (Figure 11 A). 600 mg/L
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Table 2. Degradation constants (k) for first-order substrate consumption model and specific oxygen uptake rate (SOUR) in activated sludge systems operated with (Test) and without enzymatic pre-hydrolysis (Control) in the treatment of dairy wastewaters containing different fat contents Parameter
O&G (mg/L) 400 600 800 Control 5.1 0.65 0.25 4.0 k (d-1) Test 7.8 1.61 1.16 Control 31.2 17.9 28.0 SOUR 14.4 (mg DO/gVSS d)b Test 53.7 45.6 36.9 From Rosa et al. (2006): Activated sludge systems were operated at 25oC, HRT = 20 h, average volumetric organic load = 3 kg COD/m3 d. a without fat addition; b mg DO/g VSS d = mg dissolved oxygen/g volatile suspended solids/day; Substrate consumption described according to dS = − kS ; Reactor
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Jung et al. (2002) also reported that the sludge in the bioreactor operated with previously hydrolyzed wastewater (control) also presented better sedimentability, a characteristic that is extremely important in activated sludge processes. At 800 mg O&G/L of wastewater, an accentuated increase of sludge volumetric index (SVI) in the control bioreactor (569 mL/g) becomes evident, providing further evidence that operational problems are emerging. Meanwhile, in the bioreactor fed with pre-hydrolyzed wastewater, the sludge maintained SVI values (117 mL/g) in the range of that those recommended for optimal sedimentation. Rosa et al. (2006) investigated the efficiency of the solid enzymatic preparation produced by P. restrictum in the pre-hydrolysis of dairy wastewater with high fat content (400 to 800 mg O&G/L) before biological treatment in activated sludge reactors continuously operated at room temperature (25oC). The hydrolysis of O&G present in the effluent was conducted at 30oC, for 24 h and with 0.1% (w/v) of enzymatic preparation (with lipase and protease activity of 29 U/g and 3.8 U/g). Two identical activated sludge systems operated continuously with an HRT of 20 h and 3 kg COD/m3·d, where one system was fed with crude effluent (control) and the other system was fed with enzymatically hydrolyzed effluent. Both systems produced similar COD removal efficiencies (80 to 90%); however, the O&G accumulation rate in the control bioreactor flocs was 1.7 times higher than that in the bioreactor fed with the hydrolyzed effluent. The degradation constants (k) for a first-order substrate consumption model obtained from the control and hydrolyzed bioreactors in each regime studied, considering versus not considering the biomass concentration, are presented in Table 2. The degradation constant was always higher for the bioreactor fed with effluent that had previously been hydrolyzed. The specific oxygen uptake rates (SOUR) in the different regimes studied are also presented in this Table. As the organic matter of the effluent previously hydrolyzed is presented as less complex molecules, its assimilation by the microorganisms is facilitated, which explains the higher SOUR values obtained in the hydrolyzed bioreactor in all regimes. This result is certainly related to the lesser O&G content
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accumulation on the surface and inside of the flocs in this bioreactor, which facilitates the transport and absorption of substrate and oxygen. Valladão et al. (2007) evaluated the anaerobic biodegradability of poultry industry effluents with and without enzymatic hydrolysis. Three P. restrictum-produced solid enzymatic preparation (SEP) levels (0.1, 0.5 and 1.0% (w/v)) were tested. The average lipase and protease activities of the SEP were 29 U/g and 3 U/g, respectively. The total COD removal obtained for effluent hydrolyzed or unhydrolyzed by SEP after anaerobic treatment with the different O&G concentrations tested (150, 300, 750 and 1200 mg O&G/L) is shown in Figure 12. At 150 mg O&G/L, COD removal efficiencies between the hydrolyzed and unhydrolyzed effluents did not differ substantially (67 to 76%), demonstrating that this low O&G concentration did not cause assimilation problems by the microbial consortium. However, in the absence of enzymatic hydrolysis, the increase of the O&G concentration reduced COD removal efficiency almost linearly, such that it reached only 21% with 1200 mg O&G/L. However, in tests with effluent previously hydrolyzed with 0.1% (w/v) of SEP, O&G concentration increases did not seem to affect COD removal efficiency, which ranged from 64 to 73%. At higher SEP levels (0.5 and 1.0% w/v) the COD removal efficiencies stabilized at approximately 40 and 20%, respectively, suggesting that microbial activity may be inhibited with higher SEP levels. Cirne et al. (2007) also observed the influence of lipid concentration on hydrolysis and biomethanation of a lipid-rich (triolein) model waste. For higher amounts of lipid (31%, 40% and 47%), a stronger inhibition was observed. However, the process was able to recover from the inhibition. When the effect of addition of lipase on enzymatic hydrolysis of lipids was studied, the results showed that the higher the enzyme concentration, the more accentuated was the inhibition of methane production. The authors concluded that the enzyme appeared to enhance the hydrolysis but the produced intermediates caused inhibition of the later steps in the degradation process. Since the volatile fatty acid (VFA) profiles presented similar trends for the different concentrations of lipid tested, the major obstacle to methane production was the long-chain fatty acids (LCFA) formation. The continuous addition of enzymes to the treatment systems can become costprohibitive when in situ production and/or storage are factored in. The application of an enzyme preparation for high-fat wastewater treatment would only be justified as an emergency measure, such as a sudden increase in the fat content of the bioreactor influent (fat overloads). Therefore, Damasceno et al. (2008) investigated the efficiency of a crude solid enzymatic preparation during fat shock loads in an activated sludge system operated in continuous mode. Different shock durations and intervals were evaluated, with the goal of determining the most cost-effective schedule for solid enzymatic preparation application that would not compromise biological treatment efficiency. Two identical activated sludge systems were operated for 270 days with semi-synthetic, low-fat effluent from the dairy industry as the feed material. Occasionally the bioreactors were spiked with high fat material (shock loads of 1200 mg O&G/L) and one of the bioreactors (Test) was also fed with enzymatic preparation (0.1% w/v) concurrently with shock loads additions. The Test bioreactor presented a slightly higher average COD removal efficiency when compared to the Control bioreactor (83% for Control and 90% for the Test). The effect of the enzyme addition during fat shock loads was also evaluated through the analysis of fat accumulated in the
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activated sludge flocs (Figure 13). In the Control bioreactor, the O&G content of the sludge was 1.9 to 5.1 x higher (than the Test bioreactor) for all shock load regimens. The addition of SEP during fat overloads in the reactor feed proved to be a viable alternative, maintaining efficient COD removal in the Test bioreactor over 270 days without any operational problems.
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Figure 12. Total COD removal (%) in the first batch (after 6-7 d at 35oC) with raw and pre-hydrolyzed poultry industry effluents containing different oil and grease (O&G) concentrations. Raw effluent (effluent without pre-hydrolysis); Hydro 0.1% (effluent pre-hydrolyzed with 0.1% SEP); Hydro 0.5% (effluent prehydrolyzed with 0.5% SEP); Hydro 1.0% (effluent pre-hydrolyzed with 1.0% SEP) (From Valladão et al., 2007).
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4.6. Phenol and Color Removal of Industrial Effluents Industrial activities increase significantly the water pollution due to the product purification, colorful and finishing processes. Such activities lead to high load and volumes of pollutants being known to be strongly colored, presenting large amount of suspended solids, pH broadly fluctuating, high temperature, besides high chemical oxygen demand (COD). Color is the first contaminant to be recognized in a wastewater. A very small amount of dye in water (10 – 50 mg/L) is highly visible and reduces light penetration in water systems, causing a negative effect on photosynthesis activity. The variability in wastewater composition difficults its characterization as well as the technology definition to be applied in pollution control. In general, these effluents present lower biodegradable potential, (lower BOD5/COD ratio), heavy metals and toxic compounds, including considerable flow rate which is evidenced by a higher water consumption / mass of product obtained (Correia et al., 1994). The employed dyes have been improved and/or replaced by other chemical compounds in order to be successfully applied with lower amounts of auxiliary chemicals (especially salts) permitting the environmental problems reduction associated to textile among other industries (O'Neill et al., 1999). A simple dying operation can use a series of different dyes belonging to distinct chemical classes that results in a wastewater of very complex composition. New dye molecules are developed to resist time action, sunlight exposition, water, soap, whitening agents, transpiration, etc. Antimicrobial agents are frequently used in natural fibers, like cotton, to protect them against biological degradation. The color stability and resistance difficult its removal from the wastewater since such molecules are not easily degraded under aerobic conditions prevailing in biological wastewater treatment plants (Banat et al., 1996; McMullan et al., 2001; Kunz et al., 2002, Wesenberg et al., 2003). Recently, dye removal became a research area of increasing interest, as government legislation concerning the release of contaminated effluents becomes more stringent. Most studies have been limited to the decolorization of a single dye (Kadpan et al., 2000; Knapp et al., 1997) or even to mixtures of dyes (Robinson et al., 2001; Swamy and Ramsay, 1999; Nagai et al., 2002). Although informations about the biodegradation of single dyes are useful to drive the treatment to be applied through the understanding of kinetic behavior, a biodecolorization system must sustain its ability upon exposure to real wastewater conditions. Physical and chemical methods such as adsorption (Gupta et al., 1992), coagulationflocculation (Koprivanac et al., 1993), chemical oxidation (employing oxidant agents like chlorine, H2O2 or ozone) (Ghosh et al., 1976; Kang et al., 2000; Pérez et al., 2002; Malik and Saha, 2003; Chakraborty et al., 2003), ion exchange (Correia et al., 1994), membrane separation (Porter, 1990; Buckley, 1992; Tang and Chen, 2002; Marcucci et al., 2001) and electrochemical (Kim et al., 2005) methods may be used for wastewater decolorization. Unfortunately, these methods are quite expensive and show operational problems such as development of toxic intermediates, generation of toxic sludge, lower removal efficiency, and higher specificity for a group of dyes, among others. The performance of such methods is usually related to a combination among them, including biological treatment. Nevertheless,
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this strategy leads to a more expensive treatment process (Banat et al., 1996; Kunz et al., 2002, Costa-Ferreira et al., 2003). Biodegradable tests indicate a possible reduction of BOD and COD using biological treatment. Despite of this, color removal is a slow process since most of dye molecules are not biodegradable: azo dyes are not degradable in aerobic conditions, for instance. These compounds can be, on the other hand, converted under anaerobic conditions into more toxic substances (aromatic amines obtained by the reduction of the azo bound in dye structure). The changing between aerobic - anaerobic conditions lead to good results in some situations. The dyes partial treatment is possibly obtained due to the precipitation (insoluble dyes) or adsorption in the activated sludge flocs (Porter e Snider, 1976). The microbial decolorization has received especial attention since basidiomycete fungi (white rot) have presented to be able in performing color degradation due to its unspecific enzymatic system capable to act upon lignin. Over the past decade, white rot fungi have been studied for their ability to degrade recalcitrant organo-pollutants such as polyaromatic hydrocarbons, chlorophenols, and polychlorinated biphenyls (Reddy, 1995). The low specificity of the lignin-degrading enzymes produced by these fungi suggests that they may be suitable for the degradation of textile dyes wastewater. Although dye degradation by white rot fungi has been focused on Phanerochaete chrysosporium, Pleurotus sp. and Trametes sp. have also been related to such ability (Swamy and Ramsay, 1999; Amaral et al., 2004; Levin et al., 2004; Banat et al., 1996). Trametes versicolor releases laccase as its major extracellular enzyme, a coppercontaining polyphenol oxidase (benzenediol: O2 oxidoreductase, EC 1.10.3.2) which catalyses the oxidation of phenolic compounds (Wong and Yu, 1999). Laccase can also catalyze the oxidation of organic pollutants through molecular oxygen reduction, even in the absence of hydrogen peroxide (Thurston, 1994). Moreover, environmental conditions such as pH, type of carbon source, among others seem to play an important role in decolorization by T. versicolor (Kadpan et al., 2000). Amaral et al. (2004) applied the white rot fungus Trametes versicolor for decolorization of three synthetic textile dyes (Procion Orange MX-2R (C.I. Reactive Orange 4), Remazol Red 3B (C.I. Reactive Red 23) and Remazol Black GF (C.I. Reactive Black 5)) in the presence and absence of glucose. Different initial dye concentrations were tested and approximately 97% decolorization was achieved. It was found that fungal metabolism was induced by glucose as well as pH plays an important role in the decolorization process. The absence of glucose led to higher enzymatic activities, indicating that carbon source starvation may be an interesting strategy for laccase production. However, the decolorization was more effective in the presence of glucose, reaching 97% for 50 and 100 mg dye L-1 and 87% for 300 mg dye L-1. Similar results for azo and indigo dyes were also obtained by Wong and Yu (1999). This behavior may have two explanations. One hypothesis could be that, other enzymes rather than laccase are responsible for the decolorization, as lignin and manganese peroxidase, but low activity levels of these enzymes were measured for this specific fungus. The other hypothesis could be that laccase, although being the main oxidative enzyme, requires the presence of other metabolites in the medium to act as mediators in the decolorization process. This treatment was also applied to a real wastewater from a textile industry-dyeing sector
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(containing COD = 36,000 mg/L and 43 Pt-Co units of color) led to 92% decolorization when the effluent was diluted 42 times, which corresponded to a dye concentration of 50 mg/L. The presence of other recalcitrant wastewater compounds could be responsible for the lower decolorization achieved in real systems, which caused a reduction in the performance of more than 50% when compared to the synthetic effluent. Due to a lower inhibition of the fungus enzymatic system, the dilution of these compounds might have been responsible for the improvement in the decolorization performance. Among other fungi species, Bjerkandera adusta and its isolated MnP are suitable for biodecolorization of dye baths (from black-blue through violet and red to pale yellow) containing selected azo and anthraquinone dyes to fulfill ecological standards (Mohorcic et al., 2006). Nevertheless, the employment of pure cultures in effluent decolorization is not yet a viable solution in industrial scale since contamination by other microorganisms and also loss of activity can be observed when real conditions are applied. Table 3. Potential application of peroxidases and laccases Industry activity Textile
Enzyme LiP, MnP, HRP and Lac.
Organic synthesis or general chemical industry
LiP, MnP, HRP and Lac.
Analytical/Biosensor
Lac, HRP
Biopulping Biobleaching
LiP, MnP and Lac
Food Industry
Lac, MNP, LiP
Cosmetic Oil processing
Lac HRP, MnP
Action/application Oxidize dye structures. Could be used for synthesis or color removal from effluents. Laccases are used for Denim bleaching. Oxidize a lot of organic compounds structures phenols, amines, alcohols, pahs, pcbs, agrochemicals and dioxins. Could be used for synthesis or decontamination of sites and effluents. Used for clinical exams and environmental detection. Separation and degradation of lignin in wood pulp. Replacing chlorine-based delignification Food wastewater treatment. Wine stabilization and fruit juice processing. Laccase based hair dye Sulfur oxidation. Could be an auxiliary step in biodesulfurization process
Reference Couto and Herrera, 2006. Ferreira-Leitão et al, 2003. Wesenberg et al, 2003.
Ferreira-Leitão et al, 2003. Colonna et al, 1999; Van de Velde et al, 2001; Meunier and Meunier , 1985; Kedderis and Hollenberg, 1983. Mello and Kubota, 2007. Reid, 1995. Reid and Paice, 1994. Carvalho et al, 1998. Songulahvili, 2007.
Couto and Herrera, 2006. Madeira et al, 2008.
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An alternative to the conventional treatment of industrial effluents that have been explored by different research groups is the enzyme application, proposed since 1930’s. Aitken (1993) has presented different enzyme applications of wastewater treatment which included: removal of specific compounds from complex mixtures before biological treatment; final treatment of previously biological treated wastewaters or underground waters to achieve toxicity limits; removal of persistent compounds in diluted mixtures in which the biological treatment cannot be applied; and for in-plant treatment of high concentrated pollutants and lower volumes. Nevertheless, the use of enzymes in wastewater treatment must follow some criteria before its application, since the obtained products in the enzymatic reaction must be with lower toxicity and more biodegradable, being easily removed by the subsequent treatments. Additionally, the enzyme must be able to catalyze selectively the degradation of the target compound in the real effluent, must be active and stable under treatment conditions, the reactors must be simple and the used biocatalyst must be of lower cost (Aitken et. al., 1993). The enzymes that seem to be directly related to the decolorization process are oxidases, namely laccases and peroxidases. Table 3 presents some interesting laccases and peroxidases applications as well as potential uses of these versatile biocatalysts. Ferreira-Leitão et al. (2003) compared the usefulness of the fungal lignin peroxidase (LiP) to that observed for the plant horseradish peroxidase (HRP) concerning the degradation of methylene blue (MB) and of its demethylated derivatives. The authors showed that although both enzymes are able to oxidize MB and its derivatives, HRP reactions required higher H2O2 concentrations, presented a considerably lower reaction rate, and contrary to LiP, is unable to achieve aromatic ring cleavage. Thus, lignin peroxidase would be more suitable for phenothyazine dyes degradation and color removal from waste streams. Later studies (Ferreira-Leitão et al., 2006) presented that the use of LiP for the decolorization of MB is competitive in comparison to the majority of the reported methods, regarding reaction time, range of substrate concentration and removal efficiency (Table 4). In reactions mixtures containing 50 mg/L methylene blue and carried out at 30 ºC the dye was degraded within 30 minutes. Reaction conditions were optimized concerning H2O2 addition mode to avoid the inactivation of the enzyme by H2O2 excess, the enzyme concentration to minimize cost, and the reaction temperature. Results indicated that the use of a MB: H2O2 molar ratio of 1:5 resulted efficient to remove 90% color in reactions with MB concentrations up to 50 mg/L. The enzyme stability was not affected by peroxide concentration up to 990 μM and a LiP: H2O2 molar ratio up to 1:900. The stepwise addition of the peroxide extended the possibility of using total peroxide concentrations up to 1980 μM and. LiP was stable up to 60 ºC. Recent works showed a great efficiency to degrade textile dyes, Azul Drimaren X-3LR, Azul Drimaren X – BLN, Rubinol X – 3LR, Azul Drimaren CL – R using laccase from Pleurotus ostreatus and horseradish peroxidase (data not published). The laccase applied in this study was obtained from solid residues of shimeji mushroom production. Another interesting application of peroxidase is the oxidation of sulfur compounds present in crude oil as described by Madeira et al. (2008). Its is well known that in biodesulfurization (BDS) process the low water solubility of sulfur compounds hinders its transference from the oil phase to the cells being the rate-limiting step in the metabolism of dibenzothiophenes. Thus sulfur compounds derivatives presenting higher water solubility
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could be more easily transported increasing the BDS efficiency. A stepwise evaluation was performed of the enzymatic oxidation of dibenzothiophene (DBT) by horseradish peroxidase (HRP). Reactions were carried out in monophasic organic media containing 25% (v/v) acetonitrile. The following parameters were evaluated: hydrogen peroxide concentration (0.267 to 5.34 mM); DBT: H2O2 molar ratio (1:1 to 1:20); H2O2 addition mode (single or stepwise); pH (6.0, 7.0 and 8.0) and temperature (37, 45 and 50 oC). Best results were observed in a reaction medium at pH 8.0 presenting HRP 0.06 IU/mL, DBT 0.267 mM, DBT: H2O2 molar ratio of 1:20 (stepwise peroxide addition) and incubated at 450C for 60 minutes. Under these conditions 60% of DBT was converted into DBTO (12.4%) and DBTO2 (46.4%). The observed DBT oxidation rate of 5 mmoles/mL.min.g of HRP, was 250 fold higher than the required BDS rate, of 20 µmoles/min.g of catalyst. As such a combined enzyme-microbial desulphurization process could be envisaged. Bevilaqua et al. (2002) studied the use of biological and combined biological/enzymatic treatments in phenol degradation. The studied systems were conventional batch aerobic biological followed or proceeded by enzymatic treatment. Tyrosinase extracted from the mushroom Agaricus bispora was employed. Biological treatment efficiently degraded effluents containing up to 420 mg/L of phenol, removing 97% of the COD and 99% of the phenol in 48-hour batches. Alterations in phenol concentration intake reduced treatment efficiency significantly. Enzymatic polishing of biotreated effluent removed up to 75% of the remaining phenol in a four-hour reaction with 46 U/mL of tyrosinase and 50 mg/L of chitosan (used as coagulant). Enzymatic pretreatment with 20 U/mL of tyrosinase reduced the phenol concentration by 25 % after 2 hours of reaction, although initial COD increased up to 58 %. The subsequent biological treatment of that enzymatic pretreated effluent reduced COD to 151 mgO2/L and phenol concentration to 1 mg/L in 24-hours batches. The authors concluded that enzymatic pretreatment may be a useful tool for reducing residence time in biological reactors and preventing stress caused by increasing pollutant content. Nevertheless, color generated by enzymatic reaction and COD content increases in bioreactor intake make this technology less interesting for wastewater treatment. Use of immobilized tyrosinase through development of new techniques that stabilize it and remove generated quinones may encourage combined biological/enzymatic treatments to degrade phenols, especially in emergency situations. The enzymatic treatment of phenol water-polluted with crude extract from the Agaricus bisporus mushroom and it in natura tissue is described as an efficient process by Kameda et al. (2006), being an alternative to conventional methods for specific requirements. It took three hours of treatment to remove 90 % of phenol using 200 U/mL tyrosinase activity, without pH control. The direct addition of Agaricus bisporus tissue also resulted in a phenol removal of 90 % for the same time interval, and this tissue could be reused at least three times with the same performance, proving its potential as a biocatalyst for wastewater treatment. The toxicity evaluation of the enzymatic reaction products demonstrates that they were less toxic than the original phenol wastewater.
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Table 4. Methodologies applied for methylene blue removal/degradation Methodology
Concentration Range (mg/L)
Temperatur e (0C)
pH
Time reactio n (min)
MB:H2O2 Reference
Metals supported on alumina (Cu, Co, Mn, Ni) Adsorption on kaolinite Activated carbon (groundnut shell, coconut shell, bamboo dust, rice husk, straw) Photocatalyzed Solar ligth/TiO2 Photocatalyzed UV/TiO2 Fenton Reaction
8
30
7-10
240
1:52
15
27
8.0-10.0
180
---
100-900
30
7,2
35
---
1.5-10
25-35
7.0
360
---
5-30
25
60
---
1-10
25-30
3.0; 6.7 e9 2.2-2.6
60
1:14
Salem and El-Maazawi, 2000. Gosh et al., 2002. Kannan et al., 2000.
Kuo et al., 2001. Tang et al., 2003. Dutta et al., 2001.
Cammarota and Coelho (2006) evaluated the potential application of a crude extract (non-purified) of tyrosinase from mushrooms Agaricus bispora for color removal from synthetic solutions of reactive dyes traditionally employed in textile industry (Procion Orange MX-2R, Remazol Red 3B e Remazol Black GF). Different conditions of enzyme activity: dye (type and concentration) were analyzed in phosphate buffer pH 6.0 and 30oC. After 24h of treatment, color removal of 80 %, 78 % and 56 % were achieved to Remazol Black GF, Remazol Red 3B and Procion Orange MX-2R, respectively. Soares et al. (2006) use an enzymatic cocktail consisting of laccase plus additional peroxidases, obtained from selected fungi, to decolorize textile process wastewater. The cocktail of enzymes was selected on an empirical basis based on their ability to decolorize the dye trichromy. The corresponding co-factors such as hydrogen peroxide and a redox mediator were also added to the enzymatic cocktail. The decolorization was greatest for C.I. Reactive Black 5 (91 %), followed by C.I. Reactive Red 158 (78 %), whereas C.I. Reactive Yellow 27 (17 %) was the least decolorized. Such results are in agreement with those obtained by Cammarota and Coelho (2006) and herein previously described, demonstrating that yellow dyes are more resistant to enzyme degradation. Membrane bioreactors are being increasingly used in enzymatic catalyzed transformations. However, the application of enzymatic-based treatment systems in the environmental field is rather unusual. López et al. (2002) studied the use of manganese peroxidase (MnP) from Bjerkandera sp. for continuous dye decolorization in a direct contact membrane reactor (DCMR). The system was efficient to cope with dye decolorization (95 %), operating at extremely high dye loading rate of 2.400 g/m3 per day, allowing a very fast
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decolorization in an initial period (20 min). Thereafter, a fairly constant value was reached after 40 min, with efficiency higher than 90 %, corresponding to the steady state.
5. Conclusion In environmental technology the use of enzymes cannot increase the process costs. Thus, some commercial enzymes are forbidden since the process can be economically unviable due to its high costs. Considering these factors, new technical alternatives to obtain enzymes with a reduced cost is of special attention. Solid state fermentation represents an interesting alternative to the generation of enzymes at lower costs. Agroindustrial residues like castor bean waste constitute an interesting alternative for lipase production. Moreover, since this waste contains high levels of toxicity and allergenicity, it cannot be used for others finalities as animal feed. Additionally to lipase utilization, proteases like keratinases appear as potential candidates for feather biodegradation, permitting the production of feather meal with lower energetic costs and minimizing environmental problems. Biological treatment can also be an efficient methodology to reduce toxic compounds, by the action of different enzymes, of other residues as cassava (cyanides) and coffee (tannins, caffeine and polyphenols) wastes to increase mostly proteins contents, enhancing its applicability as animal feed. Same strategy could also be applied for removal of toxic compounds in others agro-industrial wastes as olive oil ones (cake, olive mill and semi-solid residues). The employment of enzymatic preparations produced through SSF (known as solid enzymatic preparation - SEP) from low-cost industrial wastes may represent an important contribution as an adjuvant in the treatment of effluents with high O&G contents. These enzymes catalyze the hydrolysis of complex organic compounds, transforming them into substances that can be readily biodegraded by the microbial consortium present in the subsequent biological treatment. In this context, there is a wide range of scientific studies investigating enzymatic hydrolysis processes to precede traditional biological treatments with the objectives of improving the characteristics of the treated effluent and optimizing the performance of the biological treatment of interest. The development of improved techniques in this regard is especially important for the treatment of wastewater with high O&G concentrations. Concerning to phenol and color removal, developed works point the technical viability of the treatment. Nevertheless, there are still discussions about the leftovers into the effluents after these treatments; in other words, the enzyme itself and also the toxic compounds that are still soluble. So, additional treatments as chemical or biological oxidation can also be applied with the enzymatic treatment in an effluent treatment unit, aiming banish these reminders compounds. Therefore, it is possible to aggregate value to highly undesirable wastes, by the production of enzymes with greater biotechnological potential. So, low cost methodology for biocatalyst production of high aggregate value and, simultaneously, the detoxification of agro industrial wastes and/or enzymatic wastewater treatment are promising technologies that could be implemented in environmental technology.
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6. References Acuña-Arguelles, ME; Gutierrez-Rojas, M; Viniegra-González, G; Favela-Torres, E. Production and properties of three pectinolytic activities produced by Aspergillus niger in submerged and solid-state fermentation. Appl Microbiol Biotechnol, 1995 43, 808– 814. Agbor-Egbe, T; Lape Mbome, I. The effects of processing techniques in reducing cyanogen levels during the production of some Cameroonian cassava foods. J Food Composition Analysis, 2006 19, 354–363. Aguilar, CN; Augur, C; Favela-Torres, E; Viniegra-González, G. Induction and repression patterns of fungal tannase in solid-state and submerged cultures. Proc Biochem, 2001 36, 565–570. Aguilar, CN; Favela-Torres, E; Viniegra-González, G; Augur C. Culture conditions dictate protease and tannase production in submerged and solid-state cultures by Aspergillus niger Aa-20. Appl Biochem Biotechnol, 2002 102–103, 407–414. Aguilar, CN; Rodríguez, R; Gutíerrez-Sánchez, G; Augur, C; Favela-Torres, E; PradoBarragan, L.A; Ramínez-Coronel, A; Contreras-Esquivel, JC. Microbial tannases: advances and perspectives. Appl Microbiol Biotechnol, 2007 76, 47–59. Aitken, MD. Waste treatment applications of enzymes: opportunities and obstacles. Chem Eng Journal, 1993 52, B49-B58. Aliphat, S; Perie, F; Zurdo, C; Martingnon, A. Process for enzyme pretreatment of drill cutting. 1998. Patent US5725771, United States. Amaral, PFF; Fernandes, DLA; Tavares, APM; Xavier, ABMR; Cammarota, MC; Coutinho, JAP; Coelho, MAS. Decolorization of dyes from textile wastewater by Trametes versicolor. Environ Technol, 2004 25,1313-1320. Anandan, S; Anil Kumar, GK; Ghosh, J; Ramachandra, KS. Effect of different physical and chemical treatments on detoxification of ricin in castor cake. Anim Feed Sci Technol, 2005 120, 159-168. Angelidaki, I; Ahring, BK. Effects of free long-chain fatty acids on thermophilic anaerobic digestion. Appl Microbiol Biotech, 1992 37, 808-812. Aoki; N; Kawase, M. Development of high-performance thermophilic two-phase digestion process. Water Sci Technol, 1991 23, 1147-1156. Aranda, C; Robledo, A; Loera, O; Contreras-Esquivel, JC; Rodríguez, R; Aguila, CN. Fungal invertase expression in solid-state fermentation. Food Technol Biotechnol, 2006 44, 229233. Atlow, SC; Bonadonna-Aparo, L; Klibanov, AM. Dephenolization of industrial wastewaters catalyzed by polyphenol oxidase. Biotechnol Bioeng, 1983 26, 599-603. Audi, J; Belson, M; Patel, M; Shier, J; Osterloh, J. Ricin poisoning - A comprehensive review. JAMA, 2005 294, 2342-2351. Azeredo, LAI; Freire, DMG, Leite, SG; Coelho, RR. Proteases from actinomycetes interference in solid media plate assays of hyaluronidase activity. J Microbiol Methods, 2001 45, 459 – 467.
138
D. M. G. Freire, M. L. E. Gutarra, V. S. Ferreira-Leitão et al.
Azeredo, LAI; Gomes, PM; Sant´Anna Jr, GL; Castilho, LR; Freire, DMG. Production and regulation of lipase activity from Penicillium restrictum in submerged and solid-state fermentations. Curr Microbiol, 2007 54, 361-365. Azeredo, LAI; Lima, MB; Coelho, RR; Freire, DMG. A low-cost fermentation medium for thermophilic protease production by Streptomyces sp. 594 using fether meal and corn steep liquor. Curr Microbiol, 2006 20, 1-6. Bae, T-H; Han, SS; Tak, T-M. Membrane sequencing batch reactor system for the treatment of dairy industry wastewater. Process Biochem, 2003 39, 221–231. Banat, IM; Nigam, P; Slingh, D; Marchant, R. Microbial decolorization of textile-dyecontaining effluents: a review. Enzyme Microb Technol, 1996 58, 217-227. Barrett, AJ. Classification of peptidases. In: Barrett AJ, editor. Proteolytic enzymes: serine and cysteine peptidases. New York: Academic Press; 1994; 1-61. Batra, A; Saxena, RK. Potencial tannase producers from genera Aspergillus and Penicillium. Process Biochem, 2005 40, 1553-1557. Batstone, D; Keller, J; Newell, B; Newland, M. Model development and full scale validation for anaerobic treatment of protein and fat based wastewater. Water Sci Technol, 1997 36, 423-431. Battestin, V; Alves-Macedo, G. Tannase production by Paecilomyces variotii. Bioresource Technol, 2007 98,1832–1837. Becker, P; Koster, D; Popov, MN; Markossian, S; Anlianileian, G; Markl, H. The biodegradation of olive oil and the treatment of lipid-rich wool scouring wastewater under aerobic thermophilic conditions. Water Research 1999.33:653–660. Bevilaqua, JV. Utilização de tirosinase de Agaricus bispora no tratamento de efluentes contendo fenóis. M.Sc. Thesis. Rio de Janeiro: PEQ/COPPE/UFRJ; 2000; 180p. Bevilaqua, JV; Cammarota, MC; San'tanna Jr, GL; Freire, DMG. Phenol removal through combined biological and enzymatic treatments. Braz J Chem Eng, 2002 19, 151-158. Bevilaqua, JV; Freire, DMG; Santanna, LMM; Castilho, LR; Ribeiro, MCM; Gutarra, MLE; Godoy, MG. Método de tratamento de rejeito agroindustrial para destoxificação e redução da alergenicidade, com produção de lipases. 2007. Patent PI0703290-0 (Deposit), Brazil. Bietti, M; Baciocchi, E; Steenken, S. Lifetime, reduction potential and base-induced fragmentation of veratryl alcohol radical cation in aquoeous solution. Pulse radiolysis on a ligninase “mediator”. J Phy Chem A, 1998 102, 7337-7342. Bogar, B; Szakacs G; Linden JC; Pandey, A; Tengerdy. RP. Optimization of phytase production by solid substrate fermentation. J Ind Microb Biotechnol, 2003 30, 183-189. Bornscheuer, UT; Bessler, C; Srinivas, R; Krishna, SH. Optimizing lipases and related enzymes for efficient application. Trends Biotechnol, 2002 20, 433-437. Bourbonnais, R; Leech, D; Paice, MG. Electrochemical analysis of the interactions of lacase mediators with lignin model compounds. Biochim Biophys Acta, 1998 1379, 381-390. Bourbonnais, R; Paice, MG; Freiermuth, B; Bodie, E; Borneman, S. Reactivities of various mediators and laccases with kraft pulp and lignin model compounds. Appl Environ Microbiol, 1997 63, 4627-4632. Bradoo, S; Gupta, R; Saxena, R. Screening of extracellular tannase producing fungi: development of a rapid simple plate assay. J Gen Appl Microbiol, 1996 42, 325–329.
Biocatalysis in Environmental Technology
139
Brand, D; Pandey, A; Rodriguez-Leon, J; Roussos, S; Brand, I; Soccol, CR. Packed bed columm fermenter and kinetic modeling for upgrading the nutritional quality of coffee husk in solid-state fermentation. Biotechnol Prog, 2001 17, 1065-1070. Brand, D; Pandey, A; Rodriguez-Leon, J; Roussos, S; Brand, I; Soccol, CR. Relationship between coffee husk caffeine degradation and respiration of Aspergillus sp. LBPx in solid-state fermentation. Appl Biochem Biotechnol, 2002 102-103, 169-177. Brand, D; Pandey, A; Roussos, S; Soccol, CR. Biological detoxification of coffee husk by filamentous fungi using a solid-state fermentation system. Enzyme Microbial Technol, 2000 27, 127-133. Buckley, CA. Membrane technology for the treatment of dyehouse effluents. Water Sci Technol, 1992 25, 203-209. Burdsall, HH; Eslyn, WE. A new Phanerochaete with a chrysosporium imperfect state. Mycotaxon, 1974 1, 123-133. Cail, RG ; Barford, JP; Lichacz, R. Anaerobic digestion of wool scouring wastewater in a digester operated semi-continuously for biomass retention. Agric Wastes, 1986 18, 27-38. Caixeta, CET; Cammarota, MCC; Xavier, AMF. Slaughterhouse wastewater treatment: evaluation of a new three-phase separation system in a UASB reactor. Bioresour Technol, 2002 81, 61-69. Call, HP; Mück, I. Minireview: history, overview and applications of mediated ligninolytic systems, especially lacase-mediator-systems (lignozym process). J Biotechonol, 1997 53, 163-202. Cammarota, MC; Coelho, MAZ. 2006. Application of tyrosinase obtained from Agaricus bispora for color removal from textile effluents In: Third European Meeting in Oxizymes, Oeiras, Portugal. Cammarota, MC; Freire, DMG. A review on hydrolytic enzymes in the treatment of wastewater with high oil and grease content. Bioresour Technol, 2006 97, 2195-2210. Cammarota, MC; Freire, DMG; Sant'Anna Jr., GL; Russo, C; Freire, DDC; Castilho, LR. Processo de preparação e composição de preparado enzimático para tratamento de efluentes domésticos e industriais com elevado teor de gorduras, proteínas e/ou carboidratos e processo para tratamento de efluentes domésticos e industriais com elevado teor de gorduras, proteínas e/ou carboidratos (Production process and composition of an enzymatic preparation and its use for the treatment of domestic and industrial effluents of high fat, protein and/or carbohydrate content ). 2000. Patent PI0007101-3, Brazil. Cammarota, MC; Freire, DMG; Sant'Anna Jr., GL; Russo, C; Freire, DDC; Castilho, LR. Production process and composition of an enzymatic preparation and its use for the treatment of domestic and industrial effluents of high fat, protein and/or carbohydrate content. 2003. Patent PCT/BR01/00124, New Zealand. Cammarota, MC; Teixeira, GA; Freire, DMG. Enzymatic pre-hydrolysis and anaerobic degradation of wastewaters with high oil contents. Biotechnol Let, 2001 23, 1591-1595. Cardoso, PA; Mirione, E; Ernestro, M; Massaza, F; Cliffc, J; Haquee, M; Bradhurye, JH. Processing of cassava roots to remove cyanogens. J Food Compost Anal, 2005 18, 451– 460.
140
D. M. G. Freire, M. L. E. Gutarra, V. S. Ferreira-Leitão et al.
Carta-Escobar, F; Pereda-Marín, J; Álvarez-Mateos, P; Romero-Guzmán, F; Durán Barrantes, M. Aerobic purification of dairy wastewater in continuous regime. Part II: Kinetic study of the organic matter removal in two reactor configurations. Biochem Eng J, 2005 22, 117–124. Carvalho, MEA; Monteiro, MC; Bon, EPS; Sant’Anna Jr., GL. Production and characterization of Phanerochaete chrysosporium lignin peroxidases for pulp bleaching. Appl Biochem Biotechnol, 1998 70-72, 955-966. Castilho, LR; Polato, CMS; Baruque, EA; Sant’Anna Jr., GL; Freire, DMG. Economic analysis of lipase production by Penicillium restrictum in solid-state and submerged fermentations. Biochem Eng J, 2000 4, 239–247. Cavalcanti, EDC; Maciel, FM; Villeneuve, P; Lago, RCA; Machado, OL; Freire, DMG. Acetone powder from dormant seeds of Ricinus communis L.: Lipase activity and presence of toxic and allergenic compounds. Appl Biochem Biotechnol, 2007 136, 57-65. Cavaleiro, AJ; Pereira, MA; Alves, M. Enhancement of methane production from long chain fatty acid based effluents. Bioresour Technol, 2007.In press Cerda-Montalvo, ML; Contreras-Esquivel, JC; Rodriguez-Herrera, R; Aguilar, CN. Glucose diffusion on support for solid-state fermentation and its influence on tannase production profiles. Int J Chem React Eng, 2005 3, 1–10. Chadha, BS; Gulati, H; Minhas, M; Saini, HS; Singh, N. Phytase production by the thermophilic fungus Rhizomucor pusillus. World J Microbiol Biotechnol, 2004 20, 105– 109. Chakraborty, S; Purkait, MK; DasGupta, S; De, S; Basu, JK. Nanofiltration of textile plant effluent for color removal and reduction in COD. Separation Purification Technology, 2003 31, 141-151. Chao, AC; Yang, W. Biological treatment of wool scouring wastewater. J Water Pollut Control Fed, 1981 53, 311-317. Cirne, DG; Björnsson, L., Alves M, Mattiasson B. Effects of bioaugmentation by an anaerobic lipolytic bacterium on anaerobic digestion of lipid-rich waste. J Chem Technol Biotechnol, 2006 81, 1745-1752. Cirne, DG; Paloumet, X; Björnsson, L; Alves, MM; Mattiasson, B. Anaerobic digestion of lipid-rich waste—Effects of lipid concentration. Renewable Energy, 2007 32, 965-975. Collins, PJ; Kotterman, MJJ; Field, JA; Dobson ADW. Oxidation of anthracene and benzo[a]pyrene by laccases from Trametes versicolor. Appl Environ Microbiol, 1997 63, 2543-2548. Colonna, S; Gaggero, N; Richelmi, C; Pasta, P. Recent biotechnological developments of peroxidases. Trends Biotechnol, 1999 17, 163-168. Conceição, MM; Candeia, RA; Dantas, HJ; Soledade, LEB; Fernandes, VJ; Souza, AG. Rheological behavior of castor oil biodiesel. Energy Fuels, 2005 19, 2185-2188. Correia, VM; Stephenson, T; Judd, JS. Characterisation of textile wastewaters - a review. Environ Technol 1994 15, 917-929. Costa-Ferreira, M; Soares, G; Amorim, MTP. (2003) Development of an eco-sustainable biotechnical process for dye decolourisation in the textile industry. Elec J Env Agricu Food Chem. may-june. http://ejeafche.uvigo.es.
Biocatalysis in Environmental Technology
141
Couto, SR; Herrera, JLT. Industrial and biotechnological applications of laccases: a review. Biotechnol Adv, 2006 24, 500-513. Couto, SR; Sanroman, MA. Application of solid fermentation to ligninolytic enzyme production. Biochem Eng J, 2005 22, 211-219. D’Annibale, A; Sermanni, GG; Federici, F; Petruccioli, M. Olive-mill wastewaters: a promising substrate for microbial lipase production. Bioresou Technol, 2006 97, 18281833. Dalev, P; Ivanov, I; Liubomirova, A. Enzymatic modification of feather keratin hydrolysates with lysine aimed at increasing the biological value. J. Sci. Food Agri. 1997 73, 242-244. Dalmau, E; Montesinos, JL; Lotti, M; Casas, C. Effect of different carbon sources on lipase production by Candida rugosa. Enzyme Microb Technol, 2000 26, 657-663. Damasceno, FRC; Freire, DMG; Cammarota, MC. Impact of the addition of an enzyme pool on activated sludge system treating dairy wastewater under fat shock loads. J Chem Technol Biotechnol, 2008. In press. Danalewich, JR; Papagiannis, TG; Belyea, RL; Tumbleson, ME; Raskin, L. Characterization of dairy waste streams, current treatment practices, and potential for biological nutrient removal. Water Res, 1998 12, 3555-3568. De Felice, B; Pontecorvo, G; Carfagna, M. Degradation of wastewaters from olive oil mills by Yarrowia lipolytica ATCC 20255 and Pseudomonas putida. Acta Biotechnol, 2004 17, 231-239. Deschamps, AM; Otuk, G; Lebeault, JM.. Production of tannase and degradation of chestnut tannin by bacteria. J Ferm Technol, 1983 61, 55–59. Dharmsthiti, S; Kuhasuntisuk, B. Lipase from Pseudomonas aeruginosa LP602: Biochemical properties and application for wastewater treatment. J Ind Microbiol Biotechnol 1998 21, 75-80. Di Luccio, M; Capra, F; Ribeiro, NP; Vargas, GD; Freire, DMG; Oliveira, D. Effect of temperature, moisture, and carbon supplementation on lipase production by solid-state fermentation of soy cake by Penicillium simplicissimum. Appl Biochem Biotech, 2004 113-116, 173-180. Duckworth, H; Coleman, JE. Physicochemical and kinetic properties of mushroom tyrosinase. J Biol Chem, 1970 245, 1613-1625. Dunford, HB. Heme Peroxidases. New York: John Wiley and Sons Inc.; 1999. Duran, N; Esposito, E. Potential applications of oxidative enzymes and phenoloxidase-like compounds in wastewater and soil treatment: a review. Appl Cat B, 2000 28, 83-99. Duran, N; Rosa, MA; D’Annibale, AD; Gianfreda, L. Applications of laccases and tyrosinases (phenoloxidases) immobilized on different supports: a review. Enzyme Microb Technol, 2002 31, 907-931. Dutta, K; Mukhopadhyay, S; Bhattacharjee, S; Chaudhuri, B. Chemical oxidation of methylene blue using a fenton-like reaction. J Hazard Mat, 2001 B84, 57-71. Ebune, A; Al-Asheh, S; Duvnjak, Z. Production of phytase during solid-state fermentation using Aspergillus ficuum NRRL 3135 in canola meal. Bioresour Technol, 1995 53, 7–12. Elmayergi, HH; Smith, RE. Influenece of growth of Streptomyces fradiae on pepsin-HCl digestibility and methionine content on feather meal. Can J Microbiol, 1971 17, 10671072.
142
D. M. G. Freire, M. L. E. Gutarra, V. S. Ferreira-Leitão et al.
Escribano, J; Tudela, J; Gracía-Carmona, F; García-Canovas, F. A kinetic study of the suicide inactivation of an enzyme measured though coupling reactions. Biochem J, 1989 262, 597-603. Espósito, E; Canhos, VP; Durán, N. Screening of lignin-degrading fungi for removal of color from kraft mill wastewater with no additional extra carbon-source. Biotechnol Lett, 1991 13, 571-576. Faisal, M; Unno, H. Kinetic analysis of palm oil mill wastewater treatment by a modified anaerobic baffled reactor. Biochem Eng J, 2001 9, 25–31. FAO/WHO. 1991. Joint FAO/WHO Food Standards Programme. Codex Alimentarius Commission XII, Supplement 4, FAO, Rome, Italy. Ferreira-Leitão, VS; Carvalho, MEA; Bon, EPS. Lignin peroxidase efficiency for methylene blue decolouration: comparison to reported methods. Dyes and Pigments, 2007 74, 230236. Ferreira-Leitão, VS; da Silva Jr., JG; Bon, EPS. Methylene blue and azure B oxidation by horseradish peroxidase: a comparative evaluation of class II and class III peroxidises. Appl Catal B 2003 42, 213-221. Ferrer, P; Montesinos, JL; Valero, F; Solà, C. Production of native and recombinant lipases by Candida rugosa. Appl Biochem Biotechnol, 2002 95, 221-255. Freire, DMG; Teles, EMF; Bon, EPS; Sant`Anna Jr., GL. Lipase production by Penicillium restrictum in a bench scale fermenter: Media composition, agitation and aeration. Appl Biochem Biotechnol, 1997 63, 429-421. Fuchs, W; Binder, H; Mavrias, G; Braun, R. Anaerobic treatment of wastewater with high organic content using a stirred tank reactor coupled with a membrane filtration unit. Water Res, 2003 37, 902–908. Germano, S; Pandey, A; Osaku, CA; Rocha, SN; Soccol, CR. Characterization and stability of proteases from Penicillium sp. produced by solid-state fermentation. Enzyme Microb Technol, 2003 32, 246-251. Gessesse, A; Hatti-Kaul, R; Gashe, BA; Mattiasson, B. Novel alkaline proteases from alkaliphilic bacteria grown on chicken feather. Enzyme Microb Technol, 32, 519-524. Ghosh, MM; Woodard, FE; Sproul, OJ; Knowlton, PB; Guertin, PD. Treatability studies and design considerations for textile wastewater. J Water Pollut Control Fed, 1976 50, 19761985. Gianfreda, L; Xu, F; Bollag, J-M. Laccases: a useful group of oxidorredutases enzymes. Bioremediat J, 1999 3, 1-16. Giraud, E; Gosselin, L; Raimbault, M. Degradation of cassava linamarin by lactic acid bacteria. Biotechnol Lett, 1992 14, 593-598. Gombert, AK; Lopes, AP; Castilho, LR; Freire, DMG. Lipase, amylase and protease production by Penicillium restrictum in a solid-state fermentation using babassu oil cake. Process Biochem, 1999 35, 85-90. Gosh, D; Bhattacharyya, KG. Adsortion of methylene blue on kaolinite. Appl Clay Sci, 2002 20, 295-300. Gottschalk, LMF; Macedo, JMB; Bon, EPS. Lignin peroxidase production by Streptomyces viridosporus T7A - Use of corn oil as a carbon source. Appl Biochem Biotechnol, 1999 77, 771-778.
Biocatalysis in Environmental Technology
143
Grulois, P; Alric, G; Brochon, JP; Bridoux, G; Manem, J.. L'elimination des graisses par traitement biologique aérobie (The elimination of fats by aerobic biological treatment). Techniques Sciences Méthodes, 1993 5, 247-251. Gupta, GS; Shukla, SP; Prasad, G; Singh VN. China clay as an adsorbent for dyehouse wastewaters. J Environ Technol, 1992 13, 925-936. Gutarra, MLE; Cavalcanti, EDC; Freire, DMG; Castilho, LR; Sant'Anna Jr., GL. Lipase production by solid state fermentation: cultivation conditions and operation of a packedbed bioreactor. Appl Biochem Biotechnol, 2005 121, 105-116. Gutarra, MLE; Godoy, MG; Castilho, LR; Freire, DMG. Inoculum strategies for Penicillium simplicissimum lipase production by solid state fermentation using a residue of the babassu oil industry. J Chem Technol Biotechnol, 2007 82, 313-318. Hanaki, K; Matsuo, T, Nagase, M. Mechanism of inhibition caused by long-chain fatty acids in anaerobic digestion process. Biotechnol Bioeng, 1981 23, 1591-1610. Hansen, CL; Hwang, SH. Use of upflow anaerobic sludge blanket (UASB) reactor to treat whey permeate. In: Proceedings of the Sixth International Symposium on Agriculture and Food Processing Wastes. Chicago: Am Soc Ag Eng, 1990; vol. 11, 124. Hawkes, F; Donnelly, T; Anderson, G. Comparative performance of anaerobic digesters operating on ice-cream wastewater. Water Res, 1995 29, 525-533. Hernandez-Pérez, G; Goma, G; Rols, JL. Enhanced degradation of lignosulfonated compounds by Streptomyces viridosporus. Water Sci Technol, 1998 38, 289-297. Hesseltine, CW. Solid state fermentations. Biotechnol Bioeng, 1972 14, 517-532. Hofrichter, M. Review: lignin conversion by manganese peroxidase (MnP). Enzyme Microb Technol, 2002 30, 454-466. Hölker, U; Lenz, J. Solid-state fermentation - are there any biotechnological advantages? Curr Opin Microbiol, 2005 8,301–306. Jaeger, KE; Erggert, T. Lipases for biotechnology. Curr Opin Biotechnol, 2002 13, 390-397. Jaeger, KE; Reetz, MT. Microbial lipases form versatile tools for biotechnology. Trends Biotechnol, 1998 16, 396-403. Jeganathan, J; Nakhla, G; Bassi, A. Long-term performance of high-rate anaerobic reactors for the treatment of oily wastewater. Environ Sci Technol, 2006 40, 6466–6472. Jenkins, D; Richard, MG; Daigger, GT. Manual on the causes and control of activated sludge bulking and foaming, 2nd ed. Michigan: Lewis Publishers; 1993. Jordan, SN; Mullen, GJ. Enzymatic hydrolysis of organic waste materials in a solid–liquid system. Waste Manag, 2007 27, 1820–1828. Jung, F; Cammarota, MC; Freire, DMG. Impact of enzymatic pre-hydrolysis on batch activated sludge systems dealing with oily wastewaters. Biotechnol Lett, 2002 24, 17971802. Kadpan, IK; Kargi, F; McMullan, G; Marchant, R. Effect of environmental conditions on biological decolorization of textile dyestuff by C. versicolor. Enzyme Microbiol Technol, 2000 26, 381-387. Kameda, E; Langone, MAP; Coelho, MAZ. Tyrosinase extract from Agaricus bisporus mushroom and its in natura tissue for specific phenol removal. Environ Technol, 2006 27, 1209-1215.
144
D. M. G. Freire, M. L. E. Gutarra, V. S. Ferreira-Leitão et al.
Kamini, NR ; Mala, JGS ; Puvanakrishnan, R. Lipase production from Aspergillus niger by solid-state fermentation using gingelly oil cake. Process Biochem, 1998 33, 505-511. Kang, SF; Liao, CH; Po, ST. Decolorization of textile wastewater by photo-fenton oxidation technology. Chemosphere, 2000 41, 1287-1294. Kannan, N; Sundaram, MM. Kinetics and mechanism of removal of methylene blue by adsortion on various carbons: a comparative study. Dyes and Pigments, 2000 51, 25-40. Kárpáti, I; Bencze, L; Borszérki, J. New process for physicochemical pretreatment of dairy effluents with agricultural use of sludge produced. Water Sci Technol, 1995 22, 93-100. Kedderis, GL; Hollenberg, PF. Characterization of N-demethylation reactions catalyzed by horseradish peroxidase. J Biol Chem, 1983 258, 8129-8138. Kempka, AP; Lipke, NL; Pinheiro, TLF; Menoncin, S; Teichel, H; Freire, DMG; Diluccio, M; Oliveira, D. Response surface method to optimize the production and characterization of lipase from Penicillium verrucosum in solid-state fermentation. Bioprocess Biosyst Eng, 2007 31, 119 -125. Khalil, CN; Leite, L. Process for producing biodiesel fuel using triglyceride-rich oleaginous seed directly in a transesterification reaction in the presence of an alkaline alkoxide catalyst. 2006. Patent 7112229, United States. Kim, DS; Godber, JS; Kim, HR. Culture conditions for a new phytase producing fungus. Biotechnol Lett, 1999 21, 1077–1081. Kim, KGY; Lee, KB; Cho, SH; Shim, J; Moon, SH. Electroenzymatic degradation of azo dye using an immobilized peroxidase enzyme. J Hazard Mater, 2005 B126, 183–188. Kim, SH; Han, SK; Shin, HS. Two-phase anaerobic treatment system for fat-containing wastewater. J Chem Technol Biotechnol, 2004 79, 63–71. Knapp, JS; Zhang, F; Tapley, KN. Decolourisation of orange II by a wood-rotting fungus. J Chem Technol Biotechnol, 1997 69, 289-296. Koprivanac, N; Bosanac, G; Grabaric, Z; Papic, S. Treatment of wastewaters form dye industry. Environ Technol, 1993 14, 385-390. Koster, I. Abatement of long-chain fatty acid inhibition of methanogenic by calcium addition. Biol Wastes, 1987 25, 51-59. Kostinek, M; Specht, I; Edward, VA; Pinto, C; Egounlety, M; Sossa, C; Mbugua, S; Dortu, C; Thonart, P; Taljaard, L. Characterisation and biochemical properties of predominant lactic acid bacteria from fermenting cassava for selection as starter cultures. Int J Food Microbiol, 2007 114, 342–351. Krishna, C.. Solid-state fermentation systems – An overview. Crit Rev Biotechnol, 2005 25, 1-30. Kunz, A; Peralta-Zamora, P; Moraes, SG; Duran, N. Novas tendências no tratamento de efluentes têxteis. Quim Nova, 2002 25, 78-82. Kuo, WS; Ho, PH. Solar photocatalytic decolorization of methylene blue in water. Chemosphere, 2001 45, 77-83. Lagerkwist, A; Chen, H. Control of two step anaerobic degradation of municipal solid waste (MSW) by enzyme addition. Water Sci Technol, 1993 27, 47-56. Lanciotti, R; Gianotti, A; Baldi, D; Angrisani, R; Suzzi, G; Mastrocola, D; Guerzoni, ME.. Use of Yarrowia lipolytica strains for the treatment of olive mill wastewater. Bioresour Technol, 2005 96, 317-322.
Biocatalysis in Environmental Technology
145
Leal, MCMR; Cammarota, MCM; Freire, DMG; Sant’Anna Jr, GL. Hydrolytic enzymes as coadjuvants in the anaerobic treatment of dairy wastewaters. Braz J Chem Eng, 2002 19, 175-180. Leal, MCMR; Freire, DMG; Cammarota, MC; Sant’Anna Jr, GL. Effect of enzymatic hydrolysis on anaerobic treatment of dairy wastewater. Process Biochem, 2006 41, 11731178. Lei, XG; Porres, JM. Phytase enzymology, applications, and biotechnology. Biotechnol Lett, 2003 25, 1787–1794. Lekha, P; Lonsane, B. Comparative titres, location and properties of tannin acyl hydrolase produced by Aspergillus niger PKL 104 in solid-state, liquid surface and submerged fermentations. Process Biochem, 1994 29, 497–503. Lemmer, H; Baumann, M. Scum actinomycetes in sewage treatment plant. Part 2. The effect of hydrophobic substrate. Water Res, 1988 22, 761-763. Levin, L; Forchiassin, F; Viale, A. Ligninolytic enzyme production and dye decolorization by Trametes trogii: application of the Plackett-Burman experimental design to evaluate nutritional requeriments. Process Biochem, 2004 40, 1381-1387. Lima, MB; Azeredo, LAI; Coelho, RR; Freire, DMG. Thermophilic protease production by Streptomyces sp. 594 in submerged and solid state fermentations using feather meal. J Appl Microbiol, 2006 100, 641 - 647. Lin, X; Chung-Lin, L; Casale, E; Shih, JCH. Purification and characterization of a keratinases from a feather-degrading Bacillus licheniformis strain. Appl Environ Microbiol, 1992 58, 3271-3275. Linares, A; Caba, JM; Ligero, F; Rubia, T; Martínez, J. Detoxification of semisolid olive-mill wastes and pine-chip mixtures using Phanerochaete flavido-alba. Chemosphere 2003 51, 887-891. Loperena, L; Ferrari, MD; Saravia, V; Murro, D; Lima, C; Ferrando, L; Fernández, A; Lareo, C. Performance of a commercial inoculum for the aerobic biodegradation of a high fat content dairy wastewater. Bioresour Technol, 2007 98, 1045–1051. López, C; Mielgo, I; Moreira, MT; Feijoo, G; Lema, JM. Enzymatic membrane reactors for biodegradation of recalcitrant compounds. Application to dye decolourisation. J Biotechnol, 2002 99, 249-257. Madeira, LS; Ferreira-Leitão, VS; Bon, EPS. Dibenzithiphene oxidation by horseradish peroxidase in organic media: effect of the DBT:H2O2 molar ratio and H2O2 addition mode. Chemosphere, 2008 71, 189-194. Mahadik, ND; Puntambekar, US; Bastawde, KB; Khire, JM; Gokhale, DV. Production of acidic lipase by Aspergillus niger in solid state fermentation. Process Biochem, 2002 38, 715-721. Mahanta, N; Gupta, A; Khare, SK. Production of protease and lipase by solvent tolerant Pseudomonas aeruginosa PseA in solid-state fermentation using Jatropha curcas seed cake as substrate. Bioresour Technol, 2008 99, 1729–1735. Malathi, S; Chakraborty, R. Production of alkaline protease by a new Aspergillus flavus isolate under solid-substrate fermentation conditions for use as a depilations agent. Appl Environ Microbiol, 1991 57, 712-716.
146
D. M. G. Freire, M. L. E. Gutarra, V. S. Ferreira-Leitão et al.
Malik, PK; Saha, SK. Oxidation of direct dyes with hydrogen peroxide using ferrous ion as catalyst. Separation Purif Technol, 2003 31, 241-250. Marcucci, M; Nosenzo, G; Capannelli, G; Ciabatti, I; Corrieri, D; Ciardelli, G. Treatment and reuse of textile effluents based on new ultrafiltration and other membrane technologies. Desalination, 2001 138, 75-82. Marshall, KR; Harper, WJ. The treatment of wastes from dairy industry. In: Barnes D, editor. Food and Allied Industries. London: The Oitman Press; 1984; 296-376. Marten, E; Muller, RJ; Deckwer, WD. Studies on the enzymatic hydrolysis of polyesters. I. Low molecular mass model esters and aliphatic polyesters. Polym Degrad Stab, 2003 80, 485-501. Martínez, J; Borzacconi, L; Mallo, M; Galisteo, M; Viñas, M.. Treatment of slaughterhouse wastewater. Water Sci Technol, 1995 32, 99-104. Massé, L; Kennedy, KJ; Chou, S. Testing of alkaline and enzymatic hydrolysis pretreatments for oil particles in slaughterhouse wastewater. Bioresour Technol, 2001 77, 145-155. Massé, L; Massé, DI ; Kennedy, KJ.. Effect of hydrolysis pretreatment on fat degradation during anaerobic digestion of slaughterhouse wastewater. Process Biochem, 2003 38, 1365-1372. Mateos Diaz, JC; Rodríguez, JA; Roussos, S; Cordova, J; Abousalham, A; Carriere, F; Baratti, J. Lipase from the thermotolerant fungus Rhizopus homothalicus is more thermostable when produced using solid state fermentation than liquid fermentation procedures. Enzyme Microb Technol, 2006 39, 1042-1050. McGuirl, MA; Dooley, DM. Copper-containing peroxidases. Curr Opin Chem Biol, 1999 3, 138-144. McMullan, G; Meehan, C; Conneely, A; Kirby, N; Robinson, T; Nigam, P; Banat, I; Marchant, R; Smyth, WF. Microbial decolourisation and degradation of textile dyes. Appl Microbiol Biotechnol, 2001 56, 81–87. Mello, LD; Kubota, LT. Biosensors as a tool for the antioxidant status evaluation. Talanta, 2007 72, 335-358. Méndez, R; Blázquez, R; Lorenzo, F; Lema, JM. Anaerobic treatment of cheese whey: startup and operation. Water Sci Technol, 1989 21, 1857-1860. Meunier, G; Meunier, B. Peroxidase-catalyzed O-demethylathylation reactions. J Biol Chem, 1985 260, 10576-10582. Mitchell, DA; Berovic, M; Krieger, N. Overview of solid state bioprocessing. Biotechnol Annu Rev, 2002 8, 183-225. Mitchell, DA; Lonsane, BK. Definition, characteristic and potential. In: Doelle H; Mitchell DA; Rols CE. Solid substrate cultivation . London: Elsevier Applied Science; 1992; 116. Mohorcic, M; Teodorovic, S; Golob, V; Friedrich, J. Fungal and enzymatic decolourisation of artificial textile dye baths. Chemosphere, 2006 63, 1709 –1717. Mondal, K; Samanta, S; Giri, S; Pati, B. Distribution of tannic acid degrading microorganisms in the soil and comparative study of tannase from two fungal strains. Acta Microbiol Pol, 2001 50, 75–82. Mongkolthanaruk, W; Dharmsthiti, S. Biodegradation of lipid-rich wastewater by a mixed bacterial consortium. Int Biodeterior Biodegradation, 2002 50, 101-105.
Biocatalysis in Environmental Technology
147
Murthy, MVR; Karanth, NG; Rao, KSMS. Biochemical engineering aspects of solid-state fermentation. Adv Appl Microbiol, 1993 38, 99-107. Nagai, M; Sato, T; Watanabe, H; Saito, K; Kawata, M; Enei, H. Purification and characterization of an extracellular laccase from the edible mushroom Lentinula edodes and decolorization of chemically different dyes. Appl Microbiol Biotechnol, 2002 60, 327-335. Nakamoto, S; Machida, N. Phenol removal form aqueous solutions by peroxidase-catalysed reaction using aditives. Water Res, 1992 26, 49-54. Nigam, P; Singh, D. Solid-state (substrate) fermentation systems and their applications in biotechnology. J Basic Microbiol, 1994 34, 405-423. Nishitani, Y; Sasaki, E; Fujisawa, T; Osawa, R. Genotypic analyses of Lactobacilli with a range of tannase activities isolated from human feces and fermented foods. Syst Appl Microbiol, 2004 27, 109–117. Oboh, G. (2006) Nutrient enrichment of cassava peels using a mixed culture of Saccharomyces cerevisae and Lactobacillus spp. Solid media fermentation techniques. Elec J Biotechnol. January. http://www.scielo.cl/fbpe/img/ejb/v9n1/a07/ bip/ Oboh, G; Akindahunsi, AA. Biochemical changes in cassava products (flour and gari) subjected to Saccharomyces cerevisae solid media fermentation. Food Chem, 2003 82, 599–602. Oboh, G; Akindahunsi, AA; Oshodiw, AA. Nutrient and anti-nutrient contents of Aspergillus niger-fermented cassava products (flour and gari). J Food Compost Anal, 2002 15, 617– 622. Oluwole, OSA; Onabolub, AO; Mtundac, K; Mlingi, N. Characterization of cassava (Manihot esculenta Crantz) varieties in Nigeria and Tanzania, and farmers’ perception of toxicity of cassava. J Food Compost Anal, 2007 20, 559–567. Omil, F; Garrido, JM; Arrojo, B; Méndez, R.. Anaerobic filter reactor performance for the treatment of complex dairy wastewater at industrial scale. Water Res, 2003 37, 40994108. O'Neill, CO; Wawkes, FR; Hawkes, DL; Lourenço, ND; Pinheiro, HM; Delée, W. Colour in textile effluents - sources, measurement, discharge consents and simulation: a review. J Chem Technol Biotechnol, 1999 74, 1009-1018. Onifade, AA; Al-Sane, NA; Al-Musallam, AL; Zarban, S. A review: Potentials for biotechnological applications of keratin-degrading microorganisms and their enzymes for nutritional improvement of feathers and others keratins as live stock feed resources. Bioresour Technol, 1998 66, 1-11. Ozturk, I; Ubay, G; Demir, I. Hybrid upflow anaerobic sludge blanket reactor treatment of dairy effluent (HUASBR). Water Sci Technol, 1993 28, 77-81. Palma, MB; Pinto, AL; Gombert, AK; Seitz, KH; Kivatinitz, SC; Castilho, LR; Freire, DMG. Lipase production by Penicillium restrictum using solid waste of industrial babassu oil production as substrate. Appl Biochem Biotechnol, 2000 84, 1137-1145. Pandey, A. Solid-state fermentation. Biochem Eng J, 2003 13, 81-84. Pandey, A; Soccol, CR; Mitchell, D. New developments in solid state fermentation: Ibioprocesses and products. Process Biochem, 2000a 35, 1153-1169.
148
D. M. G. Freire, M. L. E. Gutarra, V. S. Ferreira-Leitão et al.
Pandey, A; Soccol, CR; Nigam, P; Brand, D; Mohan, R; Roussos, S. Biotechnological potential of coffee pulp and husk for bioprocesses. Biochem Eng J, 2000b 6, 153-162. Pandey, A; Soccol, CR; Nigam, P; Soccol, VT; Vandenberghe, LPS; Mohan, R. Biotechnological potential of agro-industrial residues. II: cassava bagasse. Bioresour Technol, 2000c 74, 81-87. Pandey, A; Szakacs, G; Soccol, CR; Rodriguez-Léon, JA; Soccol, VT. Production, purification and properties of microbial phytases. Bioresour Technol, 2001 77, 203–214. Papagianni, M; Nokes, SE; Filer, K. Production of phytase by Aspergillus niger in submerged and solid state fermentation. Process Biochem, 2000 35, 397–402. Papagianni, M; Nokes, SE; Filer, K. Submerged and solid-state phytase fermentation by Aspergillus niger: Effects of agitation and medium viscosity on phytase production, fungal morphology and inoculum performance. Food Technol Biotechnol, 2001 39, 319326. Pereira, MA; Pires, OC; Mota, M; Alves, MM. Anaerobic biodegradation of oleic and palmitic acids: evidence of mass transfer limitations caused by the accumulation of LCFA onto the anaerobic sludge. Biotechnol Bioeng, 2005 92, 15–23. Pérez, M; Torrades, F; Domènech, X; Peral, J.. Fenton and photo-Fenton oxidation of textile effluents. Water Res, 2002 36, 2703-2710. Pérez-Guerra, N; Torrado-Agrasar, A; López-Macias, C; Pastrana, L.. Main characteristics and applications of solid substrate fermentation. Electron J Environ Agric Food Chem, 2003 2, 343-350. Perle, M; Kimchie, SH; Shelef, G. Some biochemical aspects of the anaerobic degradation of dairy wastewater. Water Res, 1995 29, 1549-1554. Petruy, R; Lettinga, G. Digestion of a milk-fat emulsion. Bioresour Technol, 1997 61, 141149. Pinto, G; Leite, S; Tarzi, S; Couri, S. Selection of tannase producing Aspergillus niger strains. Braz J Microbiol, 2001 32, 24–26. Piras, A; Brigodiot, IM; Lalot, T. Degrading unsatd. polyester waste contg. crosslinking monomer, free radical catalyst and opt. filler - by suspending comminuted waste in water or alcohol and reacting in presence of enzyme. 1994. Patent WO-9426812, France. Pokorony, D; Friedrich, J; Cimerman, S. Effect of nutritional factors on lipase biosynthesis by Aspergillus niger. Biotechnol Lett, 1994 16, 363-366. Porter, JJ. Membrane filtration techniques used for recovery of dyes, chemicals and energy. Text Chem Colorist, 1990 22, 21-25. Porter, JJ; Snider, EH. Long-term biodegradability of textile chemicals. J Water Pollut Control Fed, 1976 48, 2198-2210. Radick, K. Dairy wastes. Water Environ Res, 1992 64, 417-418. Ramesh, MVL ; Lonsane, BK. Ability of a solid state fermentation technique to significantly minimize catabolic repression of α-amylase production by Bacillus licheniformis M27. Appl Microbiol Biotechnol, 1991 35, 591-593. Rana, N; Bhat, T. Effect of fermentation system on the production and properties of tannase of Aspergillus niger van Tieghem MTCC 2425. J Gen Appl Microbiol, 2005 51, 203– 212.
Biocatalysis in Environmental Technology
149
Rao, MB; Tanksale, AM; Ghatge, MS; Deshpande, VV. Molecular and biotechnological aspects of microbial proteases. Microbiol Mol Biol Rev, 1998 62, 597-635. Rao, PV; Jayaraman, K; Lakshmanan, CM. Production of lipase by Candida rugosa in solid state fermentation. 1: Determination of significant process variables. Process Biochem, 1993 28, 385-389. Rao, PVL; Jayaraj, R; Bhaskar, ASB; Kumar, O; Bhattacharya, R; Saxena, P.. Mechanism of ricin-induced apoptosis in human cervical cancer cells. Biochem Pharmacol, 2005 69, 855-865. Reddy, CA. The potential for white-rot fungi in the treatment of pollutants. Curr Opin Biotechnol, 2005 6, 320–328. Reid, ID. Biodegradation of lignin. Can J Bot, 1995 73, S1011-S1018. Reid, ID; Paice, MG. Biological bleaching of kraft pulps by white rot fungi and their enzymes. FEMS Microbiol Rev, 1994 13, 369-376. Rico, JL; García, P; Fdz-Polanco, F. Anaerobic treatment of cheese production wastewater using a UASB reactor. Bioresour Technol, 1991 37, 271-276. Riffel, A; Lucas, FS; Heeb, P; Brandelli, A. Characterization of a new keratinolytic bacterium that completely degrades native feathers. Arch Microbiol, 2003 179, 258-265. Rintala, JA; Ahring, BK. Thermophilic anaerobic digestion of source-sorted household solid waste: effects of enzyme additions. Appl Microbiol Biotechnol, 1994 40, 916-919. Rinzema, A. Anaerobic treatment of wastewater with high concentration of lipids and sulphate. Ph.D. Thesis. Wageningen: Wageningen Agricultural University; 1988. Rinzema, A; Alphenaar, A; Lettinga, G. Anaerobic digestion of long-chain fatty acids in UASB and expanded granular sludge bed reactors. Process Biochem, 1993 28, 527-537. Rinzema, A; Boone, M; van Knippenberg, K; Lettinga, G. Bactericidal effect of long chain fatty acids in anaerobic digestion. Water Environ Res, 1994 66, 40-49. Rivera –Muñoz, G; Tinoco-Valência, JR; Sànchez, S; Farrès, A. Production of microbial lipases in a solid state fermentation system. Biotechnol Lett, 1991 13, 277-280. Robinson, T; Chandran, B; Nigam, P. Studies on the production of enzymes by white-rot fungi for the decolourisation of textile dyes. Enzyme Microb Technol, 2001 29, 575-579. Rodríguez, E; Pickard, MA; Vazquez-Duhalt, R. Industrial decolorization by laccase from ligninolytic fungi. Curr Microbiol, 1999 38, 27-32. Rosa, DR; Freire, DMG; Cammarota, MC. Production and utilization of a novel solid enzymatic preparation produced by Penicillium restrictum in activated sludge systems treating wastewater with high levels of oil and grease. Environ Eng Sci, 2006 23, 814823. Roussos, S; Aquiahuati, MA; Trejo-Hernandez, M; Gaime-Perraud, I; Favela, E; Ramakrishna, M; Raimbault, M; Viniegra-Gonzalez, G. Biotechnological management of coffee pulp: isolation, screening, characterization, selection of caffeine degrading fungi and natural microflora present in coffee pulp. Appl Microbiol Biotechnol, 1995 42, 756762. Ruiz, I; Veiga, MC; de Santiago, P; Blázquez, R. Treatment of slaughterhouse wastewater in a UASB reactor and an anaerobic filter. Bioresour Technol. 1997 60, 251–258. Ryan, BJ; Carolan, N; O’Fagain, C. Horseradish peroxidase and soybean peroxidase: comparable tools for alternative niches? Trends Biotechnol, 2006 24, 355-363.
150
D. M. G. Freire, M. L. E. Gutarra, V. S. Ferreira-Leitão et al.
Saatci, Y; Arslan, EI; Konar, V. Removal of total lipids and fatty acids from sunflower oil factory effluent by UASB reactor. Bioresour Technol, 2003 87, 269–272. Saavendra, M; Benitez, E; Cifuentes, C; Nogales, R. Enzyme activities and chemical changes in wet olive cake after treatment with Pleurotus ostreatus or Eisenia fetida. Biodegradation, 2006 17, 93–102. Sabu, A; Pandey, A; Jaafar Daud, M; Szakacs, G. Tamarind seed powder and palm kernel cake: two novel agro residues for the production of tannase under solid state fermentation by Aspergillus niger ATCC 16620. Bioresour Technol, 2005 96, 1223–1228. Salem, IA; El-Maazawi, M. Kinetics and mechanism of color removal of methylene blue with hydrogen peroxide catalyzed by some supported alumina surfaces. Chemosphere 2000 41, 1173-1180. Sangali, S; Brandelli, A. Feather keratin hydrolysis by a Vibrio sp. strain kr2. J Appl Microbiol, 2000 89, 735-743. Santos, RMDB; Firmino, AAP; Sá, CM; Felix, CR. Keratinolytic activity of Aspergillus fresenius. Curr Microbiol, 1996 33, 364-370. Saxena, KL; Kaul, SN; Hasan, MZ; Gadkari, SK; Badrinath, SD. Packed bed anaerobic reactor for treatment of meat wastes. Asian Environ, 1986 8, 20-24. Sayed, SKI; van der Zanden, J; Wijffels, R; Lettinga, G. Anaerobic degradation of the various fractions of slaughterhouse wastewater. Biol Wastes, 1988 23, 117-142. Scioli, C; Vollaro, L. The use of Yarrowia lipolytica to reduce pollution in olive mill wastewaters. Water Res, 1997 31, 2520-2524. Selle, PH; Ravindran, V. Microbial phytase in poultry nutrition. Anim Feed Sci Technol, 2007 135, 1-41. Shack, PA; Shandhu, DS. Update on filament control strategies in food industries activated sludge systems. In: Proceedings of 1989 Food Processing Waste Conference, Georgia: Tech Res Inst, 1989. Sharma, R; Chisti, Y; Banerjee, UC. Production, purification, characterization, and applications of lipases. Biotechnol Adv, 2001 19, 627-662. Singh, B; Satyanarayana, T. Phytase production by a thermophilic mould Sporotrichum thermophile in solid state fermentation and its application in dephytinization of sesame oil cake. Appl Biochem Biotechnol, 2006 133, 239–250. Singh, J; Batra, N; Sobti, RC. Serine alkaline protease from a newly isolated Bacillus sp. SSR1. Process Biochem, 2001 36, 781-785. Sivalingam, G; Chattopadhyay, S; Madras, G. Solvent effects on lipase catalyzed biodegradation of poly (epsilon-caprolactone) in solution. Polym Degrad Stab, 2003 79, 413-418. Soares, GMB; Teresa, M; Amorim, P; Lageiro, M; Costa-Ferreira, M.. Pilot-scale enzymatic decolorization of industrial dyeing process wastewater. Text Res J, 2006 76, 4-11. Soares, VF; Bom, EPS; Castilho, LR; Freire, DMG. High yield Bacillus subtillis protease production by solid-state fermentation using soybean waste. Appl Biochem Biotechnol, 2005 121, 311-319. Soccol, CR; Vandenberghe, LPS. Overview of applied solid-state fermentation in Brazil. Biochem Eng J, 2003 13, 205-218.
Biocatalysis in Environmental Technology
151
Songulasvili, G; Elisashvili, V; Wasser, S; Nevo, E; Hadar, Y. Basidiomycete laccase and manganese peroxidase activity in submerged fermentation of food industry wastes. Enzyme Microb Technol, 2007 41, 57-61. Stephenson, J.A. Experience in providing high level wastewater treatment at Ontario dairy, In: Proceedings of 1989 Food Processing Waste Conference, Gerogia: Tech Res Inst, 1989. Sun, SY; Xu, Y. Solid-state fermentation for ‘whole-cell synthetic lipase’ production from Rhizopus chinensis and identification of the functional enzyme. Process Biochem, 2008 43,219–224. Swamy, J; Ramsay, JA. The evaluation of white rot fungi in the decoloration of textile dyes. Enzyme Microb Technol, 1999 24, 130–137. Takamoto, T; Shirasaka, H; Uuyama, H; Kobayashi, S. 2001. Lipase catalyzed hydrolytic degradation of polyurethane in organic solvent. Chem Lett, 6, 492-493. Tang, C; Chen, V. Nanofiltration of textile wastewater for water reuse. Desalination, 2002 143, 11-20. Tang, J; Zou, Z; Yin, J; Ye, J. Photocatalytic degradation of methylene blue on CaIn2O4 under visible light irradiation. Chem Phys Lett, 2003 382, 175-179. Tano-Debrah, K; Fukuyama, S; Otonari, N; Taniguchi, F; Ogura, M. An inoculum for the aerobic treatment of wastewaters with high concentrations of fats and oils. Bioresour Technol, 1999 69, 133-139. Thomas, A. Fats and fatty oils. Weinheim, VCH: Ullmann's Encyclopedia of Industrial Chemistry; 1987. Thorpe, SC; Kemeny, DM; Panzani, RC; Mcgurl, B; Lord, M. Allergy to castor bean .II. Identification of the major allergens in castor bean-seeds. J Allergy Clin Immunol, 1988 82, 67-72. Thurston, C F. The structure and function of fungal laccase. Microbiology, 1994 140, 19-26. Thys, RCS; Lucas, FS; Riffel, A; Heeb, P; Brandelli, A. Characterization of a protease of a feather-degrading Microbacterium species. Lett Appl Microbiol, 2004 39, 181-186. Tien, M; Kirk, TK. Lignin-degarding enzyme from hymenomycete Phanerochaete chrysosporium Burds. Science 1983 221, 661-663. Timmermans, P; Haute, A-Van, JL. Influence of the type of organisms on the biomass holdup in a fluidized-bed reactor. Appl Microbiol Biotechnol, 1993 19, 36-43. Tong, Z; Qinggxiang, Z; Hui, H; Qin, L; Yi, Z. Kinetic study on the removal of toxic phenol and chlorophenol from wastewater by horseradish peroxidase. Chemosphere, 1998 37, 1571-1577. Torkian, A; Eqbali, A; Hashemian, SJ. The effect of organic loading rate on the performance of UASB reactor treating slaughterhouse effluent. Res Conser Recycle, 2003 40, 1–11. Trudeau, F; Daigle, F; Leech, D. Reagents mediated laccases electrode for the detection of enzyme modulators. Anal Chem, 1997 69, 882-886. Tsuchiya, K; Nagashima, T; Yamamoto, Y; Gomi, K; Kitamoto, K; Kumagai, C. High level secretion of calf chymosin using a glucoamylase-prochymosin fusiongene in Aspergillus oryzae. Biosci Biotechnol Biochem, 1994 58, 895–899. Tuor, U; Winterhalter, K; Fiechter, A. Enzymes of white-rot fungi involved in lignin degradation and ecological determinants for wood decay. J Biotechnol, 1995 41, 1-17.
152
D. M. G. Freire, M. L. E. Gutarra, V. S. Ferreira-Leitão et al.
Ubalua, AO. Cassava wastes: treatment options and value addition alternatives, African J Biotechnol, 2007 6, 2065-2073. Ugwuanyi, JO; Harvey, LM; McNeil, B. Linamarase activities in Bacillus spp. responsible for thermophilic aerobic digestion of agricultural wastes for animal nutrition. Waste Manag, 2007 27, 1501–1508. Ulloa Rojas, JB; Verreth, JAJ; Amato, S; Huisman, EA. Biological treatments affect the chemical composition of coffee pulp. Bioresour Technol, 2003 89, 267–274. Uyar, F; Baysal, Z. Production and optimization of process parameters for alkaline protease production by a newly isolated Bacillus sp. under solid state fermentation. Process Biochem, 2004 39, 1893-1898. Valladão, ABG; Freire, DMG; Cammarota, MC. Enzymatic pre-hydrolysis applied to the anaerobic treatment of effluents from poultry slaughterhouses. Int Biodeterior Biodegradation, 2007 60, 219-225. van de Velde, F; van Rantwijk, F; Sheldon, RA. Improving the catalytic performance of peroxidases in organic synthesis. Trends Biotechnol, 2001 19, 73-80. van Gelder, CWG; Flurkey, WH; Wichers, HJ. Sequence and structural features of plant and fungal. Tyrosinases. Phytochemistry 1997 45, 1309-1323. Vats, P; Anerjee, UC. Production studies and catalytic properties of phytases (myoinositolhexakisphosphate phosphohydrolases): an overview. Enzyme Microb Technol, 2004 35, 3–14. Veiga, MC; Pan, M; Blázquez, R; Méndez, R; Lema, JM. A double-feed anaerobic filter for the treatment of high strength wastewaters. Biotechnol Tech, 1994 8, 77-82. Vidal, G; Carvalho, A; Méndez, R; Lema, JM. Influence of the content in fats and proteins on the anaerobic biodegradability of dairy wastewaters. Bioresour Technol, 2000 74, 231239. Villeneuve, P. Plant lipases and their applications in oils and fats modification. Eur J Lipid Sci Technol, 2003 105, 308-317. Viniegra-González, G; Favela-Torres, E; Aguilar, CN; Rómero-Gomez, SJ; Diaz-Godínez, G; Augur, C. Advantages of fungal enzyme production in solid state over liquid fermentation systems. Biochem Eng J, 2003 13, 157-167. Viraraghavan, T; Kikkeri, S. Effect of temperature on anaerobic filter treatment of dairy wastewater. Water Sci Technol, 1990 22, 191-198. Vourch, M; Balannec, B; Chaufer, B; Dorange, G. Treatment of dairy industry wastewater by reverse osmosis for water reuse. Desalination, 2008 219, 190–202. Wada, S; Ichikawa, H; Tatsumi, K. Removal of phenols form wastewater by soluble and immobilized tyrosinase. Biotechnol Bioeng, 1993 42, 854-858. Wakelin, NG; Forster, CF. An investigation into microbial removal of fats, oils and greases. Bioresour Technol, 1997 59, 37-43. Waldner, RM; Leisola, MSA; Fiechter, A. Comparison of ligninolytic activities of selected white-rot fungi. Appl Microbiol Biotechnol, 1988 29, 400-407. Walsh, J; Ross, C; Valentine, G. Food processing waste. Water Environ Res, 1994 66, 409414. Wang, Z; Banks, CJ. Evaluation of a two stage anaerobic digester for treatment of mixed abattoir wastes. Process Biochem, 2003 38, 1267–1273.
Biocatalysis in Environmental Technology
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Wesenberg, D; Kyriakides, I; Agathos, SN. White-rot fungi and their enzymes for the treatment of industrial dye effluents. Biotechnol Adv, 2003 22, 161–187. Willey, R. Fats, Oils and Greases: The minimization and treatment of wastewaters generated from oil refining and margarine production. Ecotoxicol Environ Saf, 2001 50, 127-133. Williams, CM; Richter, CS; MacKenzie Jr., JM; Shih, JCH. Isolation, identification and characterization of a feather-degrading bacterium. Appl Environ Microbiol, 1990 56, 1509-1515. Wong, Y; Yu, J. Laccase-catalyzed decolorization of synthetic dyes. Water Res, 1999 33, 3512-3520. Yamamura, S; Morita, Y; Hasan, Q; Yokoyama, KE; Tamiya, E. Keratin degradation: a cooperative action of two enzymes from Stenotrohomonas sp. Biochem Biophys Comm, 2002 294, 1138-1143. Yang, Q; Tao, L; Yang, M; Zhang, H. Effects of glucose on the decolorization of reactive black5 by yeast isolates. J Environ Sci, 2008 20, 105-108. Yang, S; Wang, J. Protease and amylase production of Streptomyces rimosus in submerged and solid state cultivations. Bot Bull Acad Sin 1999 40, 259-265.
In: Biocatalysis Research Progress Editors: F. H. Romano, A. Russo
ISBN: 978-1-60456-619-2 © 2008 Nova Science Publishers, Inc.
Chapter IV
Application of Lipases to Substances with Pharmacological Importance (Polyunsaturated Fatty Acids) Marie Zarevúcka1,* and Zdeněk Wimmer2,† 1
Institute of Organic Chemistry and Biochemistry, AS CR, Flemingovo náměstí 2, 166 10 Prague 6 – Dejvice, Czech Republic 2 Laboratory of Chemistry, Institute of Experimental Botany AS CR, Vídeňská 1083, 142 20 Prague 4 – Krč, Czech Republic
Abstract Polyunsaturated fatty acids and their derivatives are extensively studied natural compounds with high impact in human medicine. Their sources belong among the renewable resources, mostly of plant origin. Lipases are well established biocatalysts for the enantio- and regioselective formation and hydrolysis of ester bonds in a wide variety of natural and unnatural substrates. Therefore, they seemed ideally suited also for bioconversion of the plant materials, especially plant oils. The use of biocatalysts for preparation of partial acylglycerols could provide numerous advantages as compared to conventional chemical methods such as increased selectivity, higher product purity and quality, energy conservation and the elimination of application of toxic catalysts. Two general routes to desired molecules are available, in principle namely hydrolysis/alcoholysis of triacylglycerols and esterification of glycerol. The reactions can be provided under conventional conditions or in supercritical fluids. Enzymatic reactions in supercritical fluids combine the advantages of biocatalysts (substrate specificity under mild reaction conditions) and supercritical fluids (high mass* †
e-mail:
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Marie Zarevúcka and Zdeněk Wimmer transfer rate, easy separation of reaction products from the solvent, environmental benefits).
1. Introduction ω-3 Fatty acids are a type of polyunsaturated fatty acid (PUFA). PUFAs are so called because they are not 'saturated' with hydrogen atoms at multiple (poly) locations within the molecule and, as a result, contain two or more carbon-carbon double bonds. They form one of the three main classes of fatty acids, the others being saturated, in which all available hydrogen atom positions are filled, and monounsaturated, in which a single carbon-carbon double bond exists. PUFAs are subdivided into the ω-3 (n-3) series (the first double bond is 3 carbons from the ω-end carbon atom of the molecule) that are synthetically derived from linoleic acid (LA), and the ω-6 (n-6) series which are derived from α-linolenic acid (ALA), both 18 carbon atom containing fatty acids (Scheme 1). LA and ALA are termed essential fatty acids because mammalian cells are unable to synthesize these fatty acids de novo. LA can be converted sequentially via a biosynthetic pathway into other ω-6 fatty acids, the 18 carbon γ-linolenic acid (GLA), and the 20 carbon arachidonic (AA) and dihomo-γ-linolenic acids (DGLA). Similarly, ALA is converted into longer chain ω-3 fatty acids such as 20 carbon eicosapentaenoic acid (EPA) and 22 carbon docosahexaenoic acid (DHA). Increasing evidence indicates, however, that although LA and ALA can be converted into their longer chain length metabolites, the rate of conversion in humans is very slow, resulting in an estimated 2 to 10% of ALA being converted to DHA or EPA [1,2]. This suggests that a major source of the longer chain polyunsaturated fatty acid species such as EPA and DHA is likely to be dietary. Such a view is supported by data that supplementation with fish oils can markedly elevate the cellular levels of both these ω-3 PUFAs [3]. ω-3 PUFAs are of particular interest from a nutritional standpoint since the intake of these fatty acids is considered to be low in Western diets [4]. They have long been investigated for their cardioprotective and anti-inflammatory roles, which has lead to their increased use as dietary supplements [5]. A new application for ω-3 fatty acids has emerged recently, the treatment of certain forms of mental illness. Such a use is biologically plausible given that ω-3 fatty acids, in particular DHA, are abundant in the brain and are involved in, or modulate, the mechanism by which brain neurons communicate [6]. They have been shown to alter the functioning of neural systems utilizing dopamine and serotonin, both of which are thought to play an important role in mental illness and are major targets of psychoactive medications [6,7]. Furthermore, animal models of mental illness have suggested that ω-3 fatty acids can affect brain processes such as those that control mood and anxiety [8,9]. PUFAs may be directly administered as free fatty acids (FFAs), as ethyl esters or as triacylglycerols (TAG). FFAs seem to be the best absorbable pharmaceutical form for preventing cardiovascular diseases [10] and the ethyl ester is used for treating pancreatic cancer cachexia (associated chronic weight loss) [11].
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n-6 series of FAs
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n-3 series of FAs
COOH
COOH
α-linolenic acid (ALA, 18:3n-3)
linoleic acid (LA, 18:2n-6) δ6-desaturase
COOH
COOH γ-linolenic acid (GLA, 18:3n-6)
stearidonic acid (SA, 18:4n-3) elongase
COOH
COOH
dihomo- γ-linolenic acid (DGLA, 20:3n-6)
icosatetraenic acid (ITA, 20:4n-3)
1 series of prostaglandins 3 series of leukotrienes COOH
COOH
δ5-desaturase oxygenase
arachidonic acid (AA, 20:4n-6)
icosapentaenoic acid (IPA, 20:5n-3)
2 series of prostaglandins 4 series of leukotrienes
2 series of prostaglandins 4 series of leukotrienes COOH
COOH elongase adrenic acid (ADA, 22:4n-6)
docosapentaenoic acid (DPA, 22:5n-3)
Scheme 1. Metabolic pathways of polyunsaturated fatty acids.
2. PUFA Sources Natural sources of n-6 PUFAs contain variable amounts of these acids but this rarely exceeds 25% GLA and is even lower for oils other than borage oil. Oil produced from borage, evening primrose and blackcurrant seeds are rich sources of GLA and contain 1725%, 8-10% and 10-12% GLA, respectively [12-14]. Thus there has always been an interest in producing higher concentrates of GLA. Different fractionation techniques have been developed to enrich GLA from natural sources. These include urea fractionation of fatty acids
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[15-17], separation on Y-zeolite and lipase catalyzed reactions, such as selective hydrolysis of GLA-containing triacylglycerols [18], and selective esterification of GLA-containing fatty acid mixtures [19]. The only commercial sources of n-3 PUFAs (EPA and DHA) and AA are fish oils and animal viscera, respectively, e.g. AA obtained from pig liver (containing less than 0.5% AA on dry wt. basis) [20] and the fungus Morteriella alpina 1S-4 [21]. However, many microalga species have also been found to be rich in oils containing various amounts of PUFAs. Phaeodactylum tricornutum is a potential source of EPA because it is a fast growing microalga with low content in other PUFAs such as DHA and AA, which has important advantages in simplifying recovery [22]. Porphyridium cruentum UTEX 161 [22,23] is a potential source of AA as well as EPA. Marine fish oils contain substantial amounts of polyunsaturated fatty acids and are currently the major sources of EPA and DHA. Tuna oil contains approximately 5.7% EPA and 20.7% DHA [24]. Many plant sources are composed of essential long-chain fatty acids (LCFAs) such as oleic acid (C18:1), linoleic acid (C18:2) and linolenic acid (C18:3). Palm olein fractions contain a high proportion of unsaturated fatty acids, in particular C18:1 (63.1%) and C18:2 (23.9%) [25]. These LCFAs are generally recognized as useful nutrient substrates.
3. Triacylglycerol Structure Energy reserves in the form of fatty acids are in nearly all plant cells contained in a triacylglycerol molecule. The structure of triacylglycerol molecule is depicted in Fig. 1. Three fatty acids are esterified, one to each of the hydroxyl groups of a glycerol molecule, to form triacylglycerol. Since the glycerol molecule does not have rotational symmetry, all the carbon atoms are readily distinguished from each other. They are therefore classified by the sn (stereochemical numbering)-1, sn-2, and sn-3 (Figure 1) according to the recommendation of the IUPAC-IUB Commission of Biochemical Nomenclature. The plant triacylglycerols can posses a great number of different fatty acids, although eight particular ones account for some 97% of those present in commercial vegetable oils. The seed triacylglycerols are usually characterized by predominance of C18-unsaturated and polyunsaturated fatty acids, and this distinguishes them from animals fats, which are generally of a more saturated nature. The C18-unsaturated fatty acids (oleic, linoleic, and linolenic) are particularly important and govern, to a large degree, the physical properties of the oil and hence its use and commercial value. The particular fatty acids in the plant triacylglycerols are not distributed randomly between the different sn-carbon atoms. It is a general rule that saturated species of fatty acids are confined to the positions sn-1 and sn-3 with some enrichment in the first position, and that the polyunsaturated C18 fatty acids are located mainly at position sn-2 [26,27].
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sn-1
O O
H2
R C O H2
C O C C
R sn-2
H O
C O C
R
sn-3 Figure 1. Triacylglycetol structure, R = fatty acid acyl chains.
4. Enzymes Acting in Lipid Modifications Oils and fats have belonged among the most important renewable materials for the chemical industry. Industrial oleochemistry has concentrated predominantly on reactions involving carboxylic functionalities of the fatty acids. More recently, modern synthetic methods have been extensively applied to all types of natural fatty acid derivatives for the selective functionalization of their alkyl chains [28]. Application of enzymes acting in lipid modifications displays increasing role among those methods, especially when the products are considered for application in pharmacology or in food industry [29]. Industrially used hydrolysis of plant oils has been an important reaction for the production of fatty acids and glycerol as a major source of surfactants and detergents. Another process of industrial importance has been the hydrolysis of vegetable oils to enrich them with free fatty acids [30-32]. Free fatty acids, especially polyunsaturated fatty acids (PUFAs) have been of considerable pharmaceutical interest due to their biomedical properties. A replacement of the present chemical processes has been usually required, because high temperature, high pressure and presence of inorganic catalysts are involved in the hydrolysis of oil [33]. The environmentally friendly and sustainable alternatives are enzyme-catalyzed processes. The mild temperature conditions required for enzymatic hydrolysis allow energy saving and lead to a product of improved quality (color and flavor) free of traces of inorganic catalysts. Lipases and other less frequently used enzymes acting in lipid modifications can selectively lower activation energies, provide higher reaction specificities and enhance reaction rates in comparison to non-enzymatic reactions. Among those enzymes, lipases have been extensively used in triacylglycerol modification technology [34]. The enzyme-catalyzed hydrolysis can be performed in aqueous, organic or other nonconventional reaction media, e.g. supercritical fluids, ionic liquids and their combinations
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[35]. While organic solvents affect the nature of biocatalysis in several ways, including interaction with the essential hydration level of the enzyme, direct participation in the reaction mechanism, alteration of protein structure and its flexibility and alteration of the observed enzyme kinetics [36], supercritical fluids and ionic liquids represent environmentally friendly media for enzyme catalyzed processes [35,37].
4.1. Lipases Lipases (E.C.3.1.1.3) belong to the family of serine hydrolases and can be found in animals, plants, fungi and bacteria [38-41]. Because enzymes are usually named after the type of reaction they catalyze, lipases are sometimes redefined as carboxylesterases acting on long-chain acylglycerols, and are often termed triacylglycerol hydrolases. They can also catalyze the formation of acylglycerols from free fatty acids and glycerol because the reaction they catalyze is reversible (Scheme 2). In the industrially developed countries, the lipids present in the human diet consist mainly of triacylglycerols (TAGs), from 100 to about 150 g per day, i.e., 30% of each individual’s daily caloric intake. However, these TAG molecules cannot cross the intestinal barrier. A series of hydrolytic and absorption stages are necessary to produce the chemical energy resources present in the hydrocarbon chains of biologically usable TAGs. Lipases in the digestive tract play an important role in nutrition processes, both in humans and in higher animals [42]. Some lipases are able to control access to their active site. However, most of the lipases, which are used in laboratory investigations and/or in industrial production, are substrate tolerant enzymes, which accept a large variety of natural and synthetic substrates for biotransformation. Lipases do not require cofactors. They are often used in both, free or immobilized forms. They are commercially available, relatively inexpensive, and display relatively high stability. They act at the lipid–water interface and, therefore, they do not require water-soluble substrates. This function distinguishes lipases from other hydrolytic enzymes, and their efficiency in conducting transformations in organic solvents under mild conditions increases their importance as useful tools in organic synthesis [38-41,43-46]. Reactions in which lipases may be involved, both in nature and in laboratory or industrial application, are: (a) enzyme-catalyzed hydrolysis, (b) enzyme-catalyzed esterification, (c) enzyme-catalyzed transesterification by acidolysis, (d) enzyme-catalyzed transesterification by alcoholysis, (e) enzyme-catalyzed interesterification and (f) enzymecatalyzed aminolysis (Scheme 3). OCOR1 OCOR2 OCOR3
R1COOH
OH +
3 H 2O
OH
+
OH
Scheme 2. Lipase-catalyzed reversible process of a triacylglycerol hydrolysis and formation.
R2COOH R3COOH
Application of Lipases to Substances with Pharmacological Importance O
O +
R1
H2O
OR2
R1
+
R2OH
OH
O
H2O
(b)
O
OH
OR2
R3
(c) R1
OH
O +
+
R3OH
OR2
O
R1
O
O OR2
(d)
R2OH
OR3
O
+
(e)
+ R3
OR4
OR2
R3
O
R1
OR4
O +
R1
+
+ R3
O
R1
(a)
O
+
R1
R2OH
OR2
R1
O OR2
R1
+ OH
O
O R1
161
OR2
R3NH 2
+ R1
NHR3
R2OH
(f)
Scheme 3. Processes catalyzed by lipases: (a) enzyme-catalyzed hydrolysis, (b) enzyme-catalyzed esterification, (c) enzyme-catalyzed transesterification by acidolysis, (d) enzyme-catalyzed transesterification by alcoholysis, (e) enzyme-catalyzed interesterification and (f) enzyme-catalyzed aminolysis.
Lipase-catalyzed reactions are carried out under milder conditions: temperature lower than 70oC, atmospheric pressure and with higher selectivity than their chemical counter parts. In vivo, these enzymes catalyze the hydrolysis of glycerides at oil-water interfaces. In organic media with low water activity, they catalyze esterification [47] and also interesterification of TAGs via alcoholysis, acidolysis, and transesterification. A considerable number of papers has been published on lipase-catalyzed interesterifications [48-73]. Most of these works are kinetic studies on model reactions of acidolysis [51,57,58,61-63] at lab-scale, and frequently in the presence of organic solvents [51-54,61]. The bioproduction of TAGs enriched with n-3 PUFAs has also been attempted via acidolysis [64-71]. In these systems, the recovery of the modified TAG posses a separation problem. Therefore, for industrial purposes, lipasecatalyzed transesterification-ester interchange, seems to be a more adequate route than acidolysis. Several studies have been carried out, either in the presence of organic solvents [53-55] or in solvent-free media [49,55,72,73]. The fats were obtained by transesterification
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of palm oil stearin with a concentrate of TAGs enriched with n-3 PUFAs and soybean oil, catalyzed by a commercial immobilized Candida antarctica lipase (Novozyme 435) in solvent-free media. Lipases occur widely in animals, plants and microorganisms [74]. One attractive feature of lipases is their specificity with respect to the glyceride position and fatty acid type, which could seldom be constructed by chemical catalysis [75]. Although a number of lipases, especially those in microorganisms, have been well characterized, knowledge of their structure-activity relationships remains sparse, as is information about the three dimensional structures of lipases [76]. Lipase-catalyzed reactions offer several benefits over chemicallycatalyzed reactions, such as milder operation conditions [77].
4.2. Lipase Structure and Catalytic Ability In most lipases there is a mobile element, which is known as “lid”, consisting of either one or two short α-helices linked to the body of the lipase by flexible structural elements. In the open active site of lipases, the lid moves away making the binding site accessible to the substrate [78]. On the other hand, the lid needs an interface to be opened [79-83]. Three types of lipases could be identified according to their coordination-substrate site [84]. One of these types of lipases corresponds to the Rhizomucor family including Thermomyces (formerly Humicola) lanuginosa, which has an active site and a lid on the surface of the enzyme. Another type of lipases corresponds to Pseudomonas and Candida antarctica family, which has an active site and a funnel-like lid. Candida antarctica lipase B exhibits a very small lid and a funnel-like binding site. The last type corresponds to Candida rugosa family and it is characterized by the presence of an active site at the end of a tunnel containing the lid in its external part [85]. This peculiarity affects the coordination of the substrate because the reaction catalyzed by R. miehei is stimulated in position either 1 or 3 – rather than in position 2 – of the triglyceride whereas in C. rugosa the serine, which is part of the catalytic triad of the lipase, attacks all positions of the triglyceride (1 = 3 and 2) [82].
4.3. Lipoxygenases Lipoxygenases (EC 1.13.11.12) are nonheme iron dioxygenases that stereospecifically form an S-hydroperoxide under the presence of oxygen on a Z-configured double bonds (“cisdouble bond”) of a polyunsaturated fatty acid (PUFA). The Z-configured double bonds have to be separated by a single methylene junction unit to be accessible for lipoxygenases. They are very regioselective enzymes, and their actions are dependent on their own nature, i.e. lipoxygenases from different sources catalyze reactions on different double bonds located in PUFAs [29].
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4.4. Phospholipases Phospholipases involve five different enzyme types [29], PLA1 (EC 3.1.1.32), PLA2 (EC 3.1.1.4), PLB (EC 3.1.1.5), PLC (EC 3.1.4.10 and EC 3.1.4.3) and PLD (EC 3.1.4.4). PLA1 catalyzes the same reactions as 1,3-selective lipases. PLA2 selectively hydrolyzes and esterifies acyl groups at the 2-position of glycerolphospholipids. PLB displays broad substrate selectivity, hydrolyzing and forming ester bonds at the C(1) and C(2) carbon centers of glycerol. PLC is a small metalloenzyme cleaving the phosphodiester bond on the glycerol side of glycerolphospholipids. PLD hydrolyzes the diester bond of phospholipids dietal to the fatty acyl side of the diester linkage.
4.5. Lipase-catalyzed Hydrolysis under Conventional Conditions Different lipases show different specificity, which is the cleavage of various bonds, chain length and structures of the cleaved fatty acids. Bottino et al. [86] noted that eicosapentaenoic acid and docosahexaenoic acid were resistant to pancreatic lipolysis. They attributed the resistance to the location of a cis double bond near the esterified carboxyl group or to the many cis double bonds in the chain placing the terminal methyl group in a position near the ester bond, thereby causing steric hindrance. Jensen et al. [87] reported that 15 triacylglycerols containing double bonds at various positions were hydrolyzed by pancreatic lipase. The substrates containing the 2 through 7 isomers of octadecaenoic acid were resistant to pancreatic lipolysis. The discrimination was greatest against the 5 isomer in the octadecaenoic acid series, when the double bond is beyond carbon 7. Moreover, Geotrichum lipase exhibits a high degree of discrimination in favor of acids having a cis double bond at the 9 position [87]. On the other hand, Lopez-Martinez et al. [88] reported the hydrolysis of borage seed oil containing γ-linolenic acid ( 6,9,12-octadecatrienoic acid) using two kinds of microbial lipases. When borage seed oil was hydrolyzed by the Candida cylindracea lipase, γ-linolenic acid was accumulated in the non-hydrolyzed glyceride residue because of the specificity toward γ-linolenic acid exhibiting 6 unsaturation. In contrast, with the Chromobacterium viscosum lipase, there was no such accumulation of γ-linolenic acid. Using this resistance to fatty acids, when fish oil was partially hydrolyzed by the microbial lipases produced by C. cylindracea, A. niger or G. candidum, the content of n-3 polyunsaturated fatty acid in the remaining glycerides was significantly increased [89,90].
4.6. Lipase-catalyzed Reactions under Non-conventional Conditions Plant seed oils are one of the most important sustainable sources of fatty acids for present and future. The basic procedure of modification of natural triacylglycerols is hydrolysis. Plant oils also contain certain quantities of additional natural products, which are pharmacologically important: vitamins, phytosterols, phytosteroids and other compounds. These minor compounds are mainly isolated from oilseeds during the deodorization process
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within the volatile by-product known as deodorizer distillate at a quantity between 2 and 20 %, together with free fatty acids, mono-, di- and triacylglycerols [91,92]. Modern nonconventional media (supercritical fluids, ionic liquids or their combination) are nowadays applied in obtaining them by separation from plant seed oils and by subsequent enzymecatalyzed modification of natural oil [29,93-95]. 4.6.1. Properties of Supercritical Carbon Dioxide Supercritical fluids have unique properties, which can be applied to a wide range of novel chemical processes [96-101]. Among many fluids, supercritical carbon dioxide, defined as carbon dioxide above its critical point (tc = 31.3 oC, Pc = 7.4 MPa), has the added benefits of an environmentally benign nature, non-flammability, low toxicity and ready availability, and it exhibits similar properties to organic solvents. It differs from ordinary solvents in having a combination of gas-like properties (i.e., low viscosities and high diffusivities which tender them favorable for mass transfer) and liquid-like properties (i.e., solubilizing power) [102]. Moreover, these properties are tunable by the manipulation of pressure and temperature. Small variations in pressure or temperature lead to significant changes in density and density-dependent solvent properties such as dielectric constant, solubility parameters and partition coefficient. These render them more attractive as ‘green designer’ solvents and promising reaction media for environmentally benign chemical processes [102]. A scheme of the system used for extractions and enzyme-catalyzed processes operated under supercritical conditions is shown in Figure 2.
Figure 2. System for performing enzyme-catalyzed reactions in supercritical carbon dioxide: (1) compressor, (2) pressure control unit, (3) and (4) optional mixing or saturation units, (5) reactor, (6) oven, (7) rinsing valve, (8) micrometer valve, (9) trap, (10) gas meter.
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4.6.2. Properties of Ionic Liquids Ionic liquids are solutions composed entirely of ions [103-105]. They are relatively polar solvents and promote the dissolution of a vast array of pharmaceutical intermediates and final drug substance (target) molecules [106,107]. The replacement of conventional solvents in biocatalytic processes by ionic liquids could therefore overcome many of the disadvantages associated with organic solvents. The ability to modify the physico-chemical properties of these solvents by simple structural modifications to the cations or changes in the anions increases the importance of ionic liquids in organic chemistry [103,106,108-110]. 4.6.3. Lipase-catalyzed Hydrolysis The application of supercritical carbon dioxide as a solvent in enzyme-catalyzed reactions has been a matter of considerable research because of its favorable transport properties, which accelerate mass-transfer-limited enzymatic reactions [111]. The inherent gas-like low viscosities and high diffusivities of supercritical fluids increase the rates of mass transfer of substrates to the enzyme. Conversely, the liquid-like densities of supercritical fluids result in higher dissolution compared to those observed for gases. Unlike the behavior of gases and liquids, the physical properties of a supercritical fluid can be adjusted over a wide range by a relatively small change in pressure or temperature [112]. Moreover, it might be relevant to stress much lower expenses of solvent in the supercritical medium compared with those of conventional solvents [33]. Another advantage of the use of supercritical fluids (gases) as reaction media is the simple separation of oil and water in a continuously operated reactor on an industrial scale, due to their different solubility in these media. Since the solvent strength of a supercritical fluid can be varied by changing pressure and temperature, the solubility of substances can easily be regulated. It can be done continuously at the outlet of the reactor, allowing an integrated reaction and separation step, and thus simplifying the downstream processing and recycling of the solvent [33]. 4.6.3.1. Soaps One of the major chemical applications of triacylglycerols consists in a preparation of soaps. Today, preferred chemical processes operate continuously and provide quantitative yields within minutes at 100 oC, independent from the origin of the fat or oil. Energy consumption is further minimized by energy recycling [41]. At present, about two million tons of soap are produced every year by these processes. As early as in 1902, however, Connstein [113] described the application of crude lipase extracted from Ricinus seed for the preparation of pure fatty acids from tallow and plant oils within 34 hours at 30 oC in yields varying between 60 and 95%, depending on the triacylglycerol used. While this is hardly competitive to the chemical processes mainly used at present, lipase-catalyzed soap production favors are the lower depreciation costs incurred (as the equipment is limited to a stirred reactor), better soap color, and the generation of 20% glycerol water mixture as a by-product (instead of more dilute glycerol in the steam splitting process). A company in Japan has produced commercially considerable quantities of soap through Candida rugosa lipase-catalyzed hydrolysis of oils and fats in the stirred reactors at around 40 oC, in a batch mode lasting less than 48 hours [41].
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4.6.3.2. Fatty Alcohols Fatty alcohols have been another key chemical intermediate produced from triglycerides. Companies working in oleochemistry have explored lipase-catalyzed fat splitting as an alternative to generate the raw material for production of fatty alcohols. These studies revealed major drawbacks: (a) the reaction time depends strongly on the type of fat or oil used (the rate of hydrolysis at 40 oC decreases in the following order for oils from olive > soya ~ sunflower > palm > coconut > tallow), (b) the space-time yields compare poorly with those for chemical process and necessitate the construction of expensive multistage reactors, (c) expensive techniques are required for the separation of the glycerol water from the oil/lipase phase, unless supercritical carbon dioxide is used as a reaction medium, and (d) continuous use of the expensive lipase catalyst is limited by its inherent instability. A company in Japan [41] operates with a Candida rugosa lipase-catalyzed process for the preparation of highly pure unsaturated fatty acids (oleic acid, linoleic acid, α-linolenic acid, γ-linolenic acid, etc.). 4.6.3.3. Monoacylglycerols Every year, large quantities of monoacylglycerols and mixtures of mono- and diacylglycerols have been manufactured through glycerolysis of triacylglycerols. After shortpath distillation, monoacylglycerols of greater than 90% purity can be obtained. They are mild emulsifiers [HLB (hydrophilic lipophilic balance) value 3.4] permitted for use as food additives. Industrial applications include emulsification in food, cosmetics, and drug preparations [114]. Investigators deal with the preparation of monoacylglycerols through lipase catalysis, but the key problem is based on a production of mixtures of mono- and diacylglycerols. Nevertheless, the most recent investigation showed a way, how to manage the reaction to get monoacylglycerols as main products in non-catalyzed process performed in supercritical carbon dioxide [115]. In turn, there are already successful lipase-catalyzed processes available: For instance a lipase from Penicillium roquefortii is able to provide more than 90% monoacylglycerols in a one-step glycerolysis reaction [114]. 4.6.3.4. Other Applications of Lipase-hydrolysis in Supercritical Carbon Dioxide Knez et al. [116] studied the thermodynamic and kinetic properties of the immobilized lipase from Aspergillus niger (Lipolase 100T), and used this enzyme for catalysis of hydrolysis of sunflower oil in supercritical carbon dioxide. To be able to perform this process for continuous large scale application, they designed a high-pressure continuous flat-shape membrane reactor. Employing it, they achieved maximum conversion after 1 hour of the reaction at 50 oC, 90 MPa, and at a flow rate for substrates of 0.1 ml min-1. Later on they further studied this process and found the optimal concentration of lipase to be 0.07 g/ml of CO2-free reaction mixture, and the highest conversions of oleic acid (0.19 g g-1 of the oil phase) and linoleic acid (0.59 g g-1 of the oil phase) at the same conditions as before, and, in addition, at pH = 7 and an oil / buffer ratio of 1 : 1 (w/w) [117,118]. We have studied blackcurrant seed oil, which is rich in α- and γ-linolenic acid. They belong among essential polyunsaturated fatty acids [119]. The human organism is unable to synthesize these PUFAs de novo, but it is able to transform linoleic acid into PUFAs. The
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ability decreases with increasing age and with any health insufficiency. To be able to isolate α- and γ-linolenic acid from blackcurrant seed oil, we used Lipozyme, a lipase from Mucor miehei, immobilized on macroporous anionic resin [120,121]. The reaction was performed in a continuous flow reactor at 10-28 MPa and 30-50 oC with carbon dioxide saturated with oil and water (55 – 100 %) flowing up through the enzyme bed. Analysis of product composition indicated unfavorable hydrodynamics with significant mixing in the reactor when solvent interstitial velocity was lower than 4 cm min-1, while above this velocity value the flow pattern was near to plug flow. Lipase stability was very good with no activity reduction observed during a long-term experiment. The reaction rate was a function of the ratio of enzyme load to solvent volumetric flow rate. A complete hydrolysis of oil was achieved in the experiments carried out with the enzyme load of 0.8 g and CO2 flow rate of 0.4-0.9 g min1 . The effects of pressure (10-25 MPa) and temperature (30-40 oC) on the reaction rate were small, and the effects of CO2 saturation with water and of enzyme distribution in the reactor were negligible. Lipozyme displayed specificity towards linolenic acids; the release of αlinolenic acid was faster and that of γ-linolenic acid slower than the release of other constituent acids present in blackcurrant oil. The experimental system was also studied and analyzed by means of HPLC-NMR hyphenated technique [122].
4.7. Lipase-catalyzed Esterification in Organic Solvents Many lipases catalyze esterification reactions in organic solvents. TAG synthesis from DHA ethyl ester has been obtained using the lipase sp 435 from Candida antarctica immobilized on macroporous acrylic resin; more than 95% of DHA was converted to TAG by esterification at 50°C for 23 h [123]. The esterification of glycerol and n-3 PUFAs from cod liver oil was studied using three commercially available lipases: Lipozyme IM from Mucor miehei, Novozym 435 from C. antarctica and lipase PS from Pseudomonas [124]. Maximum synthesis of TAG was obtained with lipase Novozym 435, which proved to be highly active. The authors [124] studied the experimental conditions required to obtain strong esterification and a large TAG yield. Medina et al. [125] studied the production of commercially viable EPA-rich TAG from cod liver oil and the microalga P. tricornutum and TAG rich in EPA and AA from the microalga P. cruentum by esterification of these PUFAs with glycerol by lipasecatalyzed reactions.
4.8. Lipase-catalyzed Transesterification and Acidolysis under Conventional Conditions Enzyme-catalyzed transesterification reactions have been employed by many researchers for modification of fats and oils. This process involves rearrangement of fatty acid moieties within triacylglycerol molecules, with consequent improvement of the physico-chemical characteristics of fats and oils. The possible use of lipases is a subject of interest because of their performance at near-ambient conditions and ability to catalyze specific reactions. Lipases have been used as biocatalysts for the modification of fatty acid profile of vegetable
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and marine oils to produce structured lipids. Structured lipids are currently manufactured for their potential clinical benefits and for food applications. These nutraceutical lipids may provide specific fatty acids with particular properties desirable in specific disease or pathological conditions. A number of studies have focused on obtaining TAG having a combination of n-3 and n-6 PUFAs. Senanayake and Shahidi [13] tested nonspecific C. antarctica, sn-1,3 specific M. miehei, and nonspecific Pseudomonas sp. The microbial lipases were able to catalyze the incorporation of EPA and DHA in the borage and evening primrose oils to various extents. Among the lipases tested, Pseudomonas sp. gave the highest degree of EPA and DHA incorporation in both oils (32.6 and 32.3% after 36 h, in borage and evening primrose oils, respectively) followed by M. miehei and C. antarctica. Thus, the most effective lipase was selected for subsequent experiments to determine optimal acidolysis conditions. Previously, Akoh and Moussata [126] used two immobilized lipases, nonspecific SP435 from C. antarctica and sn-1,3 specific IM60 from Rhizomucor miehei as biocatalysts for the restructuring of borage oil to incorporate EPA and capric acid (10:0) with free fatty acids as acyl donors. They obtained a higher incorporation of EPA (10.2%) and capric acid (26.3%) using IM60 lipase, compared to 8.8 and 15.5%, respectively, when SP435 lipase was used. The ability of immobilized lipases IM60 from M. miehei and SP435 from C. antarctica to modify the fatty acid composition of selected vegetable oils (canola, peanut, and soybean oils) by incorporation of n-3 polyunsaturated fatty acids into the vegetable oils was studied [67]. These authors used free acid and ethyl esters of EPA and DHA as acyl donors. Using free EPA as acyl donor, IM60 gave higher incorporation of EPA than SP435. However, when ethyl esters of EPA and DHA were used as acyl donors, SP435 gave higher incorporation of EPA and DHA than IM60.
4.9. Structured Lipids Structured lipids (SLs) are triacylglycerols or phospholipids in which fatty acids have been placed in specific locations in the glycerol backbone and are produced using a chemical or enzymatic process. SLs are new generation fats or oils with medical, nutraceutical and food applications. Much attention is being paid to SLs due to their potential biological functions, industrial applications and nutritional perspectives. Lipids can be restructured to meet essential fatty acid requirements or to incorporate specific fatty acids of interest into specific locations of the glycerol backbone of TAG. SL may offer the most efficient means of delivering target fatty acids for nutritive or therapeutic purposes as well as to alleviate specific disease and metabolic conditions. The constituent fatty acids and their locations in the glycerol backbone determine the functional and physical features, the metabolic fate, and the health benefits of SLs. Therefore, designing SLs with selected fatty acids at specific locations in the TAGs for medicinal applications has attracted much attention. The position of FA in the TAG molecules (sn-1, sn-2, and sn-3) will have a significant impact on their metabolism in the body. In general, FAs at the terminal positions of TAG (sn-1 and sn-3) are hydrolyzed by pancreatic lipase and absorbed while those at the middle position of TAG (sn-2) remain unchanged and are used in
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the synthesis of new TAG. For example, it may be desirable to develop a SL containing polyunsaturated fatty acids at the sn-2 position with medium-chain fatty acids (MCFAs) at the sn-1,3 positions for patients with mal-digestion as well as cystic fibrosis. A SL containing MCFA and linoleic acid is more efficient in cystic fibrosis patients than safflower oil, which has about twice as much of linoleic acid [127]. SLs have many industrial applications and have recently attracted the attention of food manufacturers for production of low-caloric lipids that are characterized by a mixture of short-chain fatty acids (SCFAs) and/or MCFAs and LCFAs in the same glycerol moiety. Increasing interest in such products stems from the fact that they contain 5-7 kcal g-1 energy compared to 9 kcal g-1 for usual fats and oils; this is because of the lower caloric content of SCFA or MCFA compared to LCFA. Reduced-calorie specialty lipids are intended for use in baking chips, dips, coatings, bakery and dairy products, or as a cocoa butter equivalent. Over the past two decades several research groups have successfully incorporated MCFAs (caprylic acid or capric acid) into fish and marine oils containing PUFAs [70,126131] into borage oil rich in γ-linolenic acid [126,128,131] and into single-cell oils [132-135]. SLs may be produced by incorporation of selected fatty acids into an oil. The degree of reactivity of different fatty acids may vary in different systems due to factors such as the lipase type, water activity, and other conditions [136]. Many lipases have been shown to be more selective toward C18 FA with higher degrees of unsaturation in esterification and interesterification reactions (C18:0 < C18:1 < C18:2) [137]. Yang et al. [136] compared incorporation of linoleic and conjugated linoleic (CLA) acids into tristearin (SSS) in a solvent-free system at 60oC using 5% Lipozyme RM IM from Rhizomucor miehei. Incorporation of LA into SSS was higher than that of CLA and suggested that the rigidity of CLA might have been responsible for this observation [136]. Tsuzuki [138] screened ten lipases for their ability to catalyze acidolysis of triolein with SCFAs (C2:0, C3:0, and C4:0) in organic solvents. Lipase from Aspergillus oryzae afforded the highest yields of products in the reaction of triolein with C2:0, C3:0, and C4:0. The results of the study indicated that as the chain length decreased, the degree of incorporation of SCFAs into triolein increased. Paez et al. [139] reported that incorporation of caprylic acid (C8:0) into triolein was favored compared with that of oleic acid. Again chain length of the FA might play a role in the observed trends. The synthesis of a modified oil via acidolysis of trilinolein (tri C18:2) with C8:0, using Lipozyme IM-60 as a biocatalyst has been reported [140]. Lipozyme IM-60 was found to be more effective for incorporation of a LCFA than a MCFA. A synthesis of SL by interesterification of trilinolein and tricaproin with sn-1,3-specific (IM 60) and nonspecific (SP 435) lipases was reported [141]. In general, it was found incorporation of selected LCFAs into TAGs (e.g. tristearin or triolein) may be affected by many factors, including chain length, number of double bonds, and the location and geometry of the double bonds as well as reaction conditions and reactivity and specificity of lipases employed. LA was more reactive than CLA due to the rigidity of the latter and/or specificity of the enzymes [142]. EPA was more reactive than DHA, due to the structural differences between the two (number of double bonds, chain length) [143]. Lipases, Novozyme-435 enzyme from C. antarctica and AY-30 from Candida rugosa, might be considered as promising biocatalysts for acidolysis of tristearin and selected LCFAs [144]. The high percent incorporation of FA into tristearin using lipase from C. antarctica or C. rugosa might be due to the experimental
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conditions employed in the study which were suitable for these two enzymes [143]. Hamam and Shahidi [143] reported the acidolysis of tristearin) and triolein [142] with LCFAs. In the study, they examined the effect of the chain length, number of double bonds, and the location and geometry of double bonds on the incorporation of selected FA into tri C18:2 and tri C18:3. Several studies have dealt with the synthesis of SL enriched in PUFAs using a one-step lipase catalyzed synthesis [135,145,146]. However, this synthesis required long reaction times and achieved only low yields of the desired products. The two-step reaction was recently suggested as an effective method for the SL synthesis resulting in higher yields and purer products than the conventional one-step reaction [147,148]. In the first-step, 2-MAG are produced by alcoholysis of a triglyceride with ethanol using a 1,3-regiospecific lipase. The 2-MAG thus obtained can subsequently be esterified with suitable fatty acids.
4.10. Lipase-catalyzed Esterification in Supercritical Carbon Dioxide Lipase-catalyzed production of various types of esters has increased tremendously within the recent years. Esters are mainly present in oils, fats and natural polymers, and they are useful intermediates or target products in chemical industry. Japanese authors [149, 150] described acylglycerol and aliphatic ester synthesis by lipases, and proved the occurrence of reverse reaction. Lipase from Aspergilus niger, Rhizopus delemar, Geotrichum candidum and Penicillium cyclopium were found to synthesize esters of oleic acid and a number of primary alcohols. From these lipases, only that from G. candidum was able to synthesize esters of secondary alcohols. Lipase-catalyzed synthesis of esters can be achieved either by reaction between free fatty acid and alcohol or by ester exchange or transesterification, which includes alcoholysis, acidolysis and interesterification [43]. Lipase-catalyzed esterification reactions have been applied in a production of different types of important esters: (a) esters of shortchain alcohols and short-chain fatty acids (aroma compounds; [43]), (b) esters of short-chain alcohols and long-chain fatty acids (oleochemicals, e.g. lubricants, diesel fuel, and antistatic reagents; [43]), (c) esters of polyhydroxy alcohols (glycerol, alcoholic sugars, carbohydrates etc.) and long-chain fatty acids, which are generally called emulsifiers or surfactants (important products for application in food and pharmaceutical industries; [43]). Lipases have often been used for treatment and modification of oils and fats [43]. Many lipases exhibit sn-1,3 specificity and may be used for regioselective (inter)esterification of natural triacylglycerols. The acyl migration observed from the sn-2 to the sn-1 or sn-3 positions must be suppressed or eliminated, wherever possible. By partial or total exchange of fatty acyls in triacylglycerols of given origin it is possible to modify physico-chemical properties and also nutritional value of the starting natural oil or fat [43]. From this point of view, three types of triacylglycerol modification are commercially and pharmacologically important: (a) production of cocoa butter equivalents, (b) production of fats with improved spreadability and (c) production of highly digestive triacylglycerols [41, 43].
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4.10.1. Cocoa Butter Equivalent The main producers of cocoa butter are tropical countries, mainly Kenya and Malaysia. Since the melting point of cocoa butter is around human body temperature (37 oC), it is well suited as a matrix for suppositories. The main application, however, is in the production of chocolates, where the rapid melting conveys a desirable feeling. The predominant triacylglycerols of cocoa butter are compounds bearing oleic acid in the sn-2 position and stearic and palmitic acids in the sn-1 and sn-3 positions; these triacylglycerols are generally known as SOS and SOP. Cocoa butter equivalents can be prepared either chemically or by lipase catalysis through the interesterification of suitable natural triacylglycerols, for example, the middle fraction of palm oil (POP) or sunflower oil with a high content of oleic acid (high-oleic sunflower oil; OOO) with stearic acid or tristearin (SSS) [41,151]. The primary hydroxyl groups of glycerol in positions sn-1 and sn-3 are more reactive than the secondary hydroxyl group in the sn-2 position, and triacylglycerols of the type SOP or SOS are predominantly formed. 4.10.2. Fats with Improved Spreadability The melting point of any oil can be modulated by the degree of catalytic hydrogenation of double bonds in unsaturated fatty acids. This is done on a large scale for the preparation of margarines and shortenings from plant oils. Alternatively, the desired melting point can be achieved through interesterification of suitable triglyceride mixtures with the use of sn-1,3 specific lipases [41]. 4.10.3. Highly Digestive Triacylglycerols The absorption of triacylglycerols from the small intestine strongly depends on their structure. Triacylglycerols containing palmitic acid are well adsorbed only when this fatty acid is located in the sn-2 position, like in human milk. A commercial product of this type (OPO) is a diet additive for premature infants. It is prepared by interesterification of tripalmitin with oleic acid with use of immobilized Rhizomucor miehei lipase. Alternatively, triacylglycerols of the MLM type, with long-chain saturated or unsaturated fatty acids (L) in the sn-2 position and medium-chain fatty acids (M) in the positions sn-1 and sn-3, provide a rapid delivery of energy through enhanced hydrolysis and resorption; pancreatic lipase preferentially hydrolyzes medium-chain triacylglycerols, and the resulting monoacylglycerols are efficiently absorbed from the large intestine. Several products of this type are commercially available, which contain polyunsaturated (essential) fatty acids. They have shown beneficial effects against cardiovascular and inflammatory diseases. Functional triacylglycerols of this composition are preferentially prepared by means of sn-1,3 specific lipases during an interesterification process starting from highly unsaturated triacylglycerols [41]. 4.10.4. Other Types of Lipase-catalyzed Reactions in Supercritical Carbon Dioxide The utilization of supercritical carbon dioxide as a reaction medium confers many advantages, among which are an environmental compatibility, zero chemical residues in the synthesized product, and considerable processing flexibility [152]. When lipase is used for
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catalyzing a synthetic procedure in supercritical carbon dioxide [153], the process is particularly applicable to producing additives that can be incorporated directly into food formulations. Lipase-catalyzed reactions in supercritical carbon dioxide have already been reported [43,154-158]. Among others, a synthesis of simple esters [159] was reported, transesterifications to make methyl esters [153] were conducted, a glycerolysis process [153] was studied, and randomization of fats/oils [153] in supercritical carbon dioxide was performed, and all those processes used Candida antarctica lipase [160], commercially known as Novozym 435. High quantitative yields when performing such transesterifications to make methyl esters have permitted application of the lipase reaction in supercritical carbon dioxide as an analytical method for quantifying fat levels in food products which are required under new food nutritional labeling guidelines [161]. Recently, a Novozym 435-catalyzed transesterification has been utilized as the initial step in a two-stage synthesis conducted under supercritical fluid conditions to produce fatty alcohols directly from vegetable oils [162]. An interesting study has been performed with lipase-catalyzed esterification of stearic acid with ethanol, and subsequent hydrolysis of ethyl stearate under the conditions set near to the critical point in supercritical carbon dioxide [163]. Pressure ranged from 6 to 20 MPa, and the temperature ranged from 35 to 60 oC. The authors [163] observed the esterification rate of stearic acid increasing near the critical point and keeping the increase steady with increasing the pressure, reflecting the increasing solubility of stearic acid in supercritical medium. The hydrolysis rate of ethyl stearate showed its maximum at a pressure near the critical point, and it was dependent on the initial concentration of ethyl stearate in the system, i.e. dependent on the fact if the quantity of the ester was completely or partly soluble in supercritical carbon dioxide. When the reaction was performed with an initial ester concentration below the solubility limit in supercritical carbon dioxide, the authors observed the maximum pressure shifted along the extended line of the gas-liquid equilibrium in the supercritical region in the pressure-temperature phase plan. This finding seems to be related to the singular behavior of some properties of supercritical carbon dioxide along this line reported in the literature [164]. Myristic acid was esterified by ethanol using a hog pancreas lipase in supercritical carbon dioxide in 37 % yield, while a similar enzyme-catalyzed reaction performed in acetonitrile yielded only 4 % of the required product [165]. Immobilized lipases from Humicola lanuginosa and Candida antarctica (lipase B) were employed for modification (alcoholysis and glyceride synthesis) of cod liver oil with ethanol in supercritical carbon dioxide [166]; fish oil is rich in important polyunsaturated fatty acids, like many plant and seed oils, and unlike animal fats. Long-chain fatty acid esters with fatty alcohols (long-chain 1-alkanols) are useful functional molecules in pharmaceutical, cosmetic and lubricant industry. Knez et al. [167,168] used lipase from Rhizomucor miehei (Lipozyme RM IM) as catalyst for production of 1-octyl oleate in supercritical carbon dioxide, and studied the influence of parameter changes to the conversion rate in bench-scale packed-bed reactor. The authors got 93 % maximum yield of the product at a substrate flow rate of 18 ml h-1 and the CO2 flow rate of 210 ml h-1, and the immobilized enzyme was able to act as catalyst up to 50 days. They proved a possibility of using such a system for continuous production process.
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Modification of castor oil in supercritical carbon dioxide may be an example of lipasecatalyzed interesterification between castor oil triacylglycerols and methyl oleate. A conversion of about 90 % was achieved after 5 hours of reaction time using immobilized C. antarctica lipase B as the catalyst [169]. Even naturally occurring aliphatic amides, like capsaicin (8-methyl-N-vanillyl-6nonenamide), a compound found in the red pepper seeds, which promotes blood circulation, can be compounds of priority interest for pharmacology. This compound is relatively hardly available in broad scale and, therefore, a search for its bioanalogs has been performed. Palmitoyl vanillylamide, one of those analogs, can be synthesized through amidation by immobilized Mucor miehei lipase in supercritical carbon dioxide under optimized conditions (50 oC, 17 MPa and pH 8 kept for 8 hours) from vanillylamide hydrochloride and palmitic anhydride at a molar ratio 5 / 15, and enzyme concentration of 0.5 % (w/w) [170]. The course of the lipase-catalyzed reactions in supercritical carbon dioxide is always affected by the presence of moisture. Japanese authors [171] studied continuous acidolysis of triolein and stearic acid using the moist immobilized lipase in supercritical carbon dioxide as a process for large scale synthesis. They found optimum operation conditions at 50 oC, 16.9 MPa and adsorbed water concentration of 2 % (w/w). The production rate was found to be about 0.03 mmol h-1 per each 1 g of the immobilized enzyme employed. 4.10.5. Separation and Lipase-catalyzed Modifications of Phytosterols and Related Minor Compounds A need has existed for developing new processing methods of facilitating plant oils extraction and refining while sustaining the nutritional components naturally present in edible oils and reducing the adverse impact of oil processing on environment [172]. Therefore, supercritical carbon dioxide extraction and fractionation techniques have been positively examined as alternative methods of obtaining plant and seed oils with high purity and quality [172,173]. Moreover, minor components of plant and seed oils, such as vitamins, phytosterols and isoprenes (e.g. squalene) can be separated. One of the most known and most commercialized oils is Canadian canola oil. Value-added processing of canola oil deodorizer distillate has benefited the canola oil processing industry in Canada considerably. Canola is one the leading plant oils and its deodorizer distillate is a concentrated mixture of phytosterols and tocopherols [174]. Synthesis of esters derived from phytosterols or other steroid alcohols and fatty acids is of great importance, due to their recent recognition and application in the food industry as cholesterol-lowering agents. Several enzymes were screened as catalysts, and optimal conditions were determined for the reaction between various fatty acids and sterols/phytosterols (e.g., cholesterol, sitostanol etc.) in supercritical carbon dioxide [153,175]. Using an analytical supercritical fluid extraction unit, the lipase derived from Burkholderia cepacia, Chirazyme L-1, was determined to be the most selective for facilitating the desired reactions [176]. Fatty acids C8–C18, a pressure range of 20.7 – 31 MPa, a temperature range of 40 – 60 °C, along with variable flow rates, and initial static hold times were used to evaluated the feasibility of the above reaction. The yield of the cholesterol esters, as measured by supercritical fluid chromatography, ranged from 90 % for caprylic acid to 99 % for palmitic acid, while the corresponding reaction between sitostanol and the same
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fatty acids produced yields of 92 % for caprylic acid and 99 % for palmitic acid, respectively [153]. The extraction apparatus was modified to provide a continuous flow of the reagent fatty acid and phytosterol or other steroid alcohols over the enzyme bed, thereby allowing continuous production of the desired esters, which averaged a 99% yield under optimal conditions [153]. 4.10.6. Lipase-catalyzed Reactions in Ionic Liquid / Supercritical Carbon Dioxide Biphasic Systems Spanish and French authors [177,178] described a synthesis of butyl esters (propanoate, laurate etc.) in a recirculating bioreactor in room temperature using ionic liquid / supercritical carbon dioxide biphasic systems at 50 oC and 8 MPa. In these systems, α-alumina microporous membranes with immobilized Candida antarctica lipase B were coated with four different ionic liquids based on 1-n-alkyl-3-imidazolium cations and hexafluorophosphate and bis{(trifluoromethyl)sulfonyl}imide anions. Selectivity increased up to > 99.5 % when the ionic liquid / supercritical carbon dioxide biphasic system was used rather than supercritical carbon dioxide alone (at room temperature). It was observed that the activity in room temperature ionic liquid / supercritical carbon dioxide biphasic systems depends on the effect of the ionic liquid media on the enzyme and the diffusion limitations across the ionic liquid layer around the biocatalyst.
5. Importance of the Reaction Media for Sustainability Extraction of compounds from natural sources has been the most widely studied application of supercritical fluids with several hundreds of published scientific papers (for review, see e.g. Reverchon and De Marco [37]. Supercritical fluid extraction has immediate advantages over traditional extraction techniques: it is a flexible process due to the possibility of continuous modulation of the solvent power/selectivity of the supercritical fluid, allows the elimination of polluting organic solvents and of the expensive post-processing of the extracts for solvent elimination. Supercritical carbon dioxide, a ‘green’ and sustainable medium, is nowadays the most frequently used supercritical fluid for productions in nutraceutic, cosmetic and pharmaceutical industry. Supercritical fluid extractions, fractionations of the extracts, and their enzyme-catalyzed modifications have been developed and have expanded tremendously in many fields of products. The processes have nowadays been developed for extraction and modification of essential and vegetable oil production, nutraceutic, cosmetic and pharmaceutical industries. This approach is also scientifically challenging but also necessary to find applications that are industrially competitive with the traditional processes based on cheaper technologies and plants. In some cases, the extraction problems proved to be very complex and as a consequence more evolved process schemes (multistep extractions, continuous solid processing, multistage separations, co-solvent or ionic liquid applications) have been adopted to overcome these problems. However, a large quantity of work is still required. Nevertheless, numbers of reviews have recently appeared which have shown perspectives of
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application of supercritical fluid technology in many areas of research and industrial production [37,43-45, 106,107,179-181].
Acknowledgment The author (MZ) thanks to the project of the Institute of Organic Chemistry and Biochemistry Z40550506. The author (ZW) thanks the Ministry of Education for the grant 2B06024 (SUPRAFYT), a part of the National Research Program II, for funding, and Mrs. M. Wimmerová for helpful discussion.
References [1]
Emken, E.A.; Adlof, R.O.; Gulley, R.M. Dietary linoleic acid influences desaturation and acylation of deuterium-labeled linoleic and linolenic acids in young adult males. Biochim. Biophys. Acta, 1994 1213, 277-288. [2] Holub, D.J.; Holub, B.J. Omega-3 fatty acids from fish oils and cardiovascular disease. Mol. Cell Biochem., 2004 263, 217-225. [3] Peet, M.; Brind, J.; Ramchand, C.N.; Shah, S.; Vankar, G.K. Two double blind placebo controlled pilot studies of eicosapentaenoic acid in the treatment of schizophrenia. Schizophr. Res., 2001 49, 243-251. [4] Simopoulos, A.P. Omega-3 fatty acids in health and disease and in growth and development. Am. J. Clin. Nutr., 1991 54, 438-63. [5] von Schacky, C. Omega-3 fatty acids and cardiovascular disease. Curr. Opin. Clin. Nutr. Metab. Care, 2004 7, 31-136. [6] Stahl, S.M. Essential psychopharmacology. Neuroscientific basis and practical applications 2nd edition. Cambridge, UK: Cambridge University Press, UK; 2000. [7] Glen, A.I.M.; Ross, B.M. Eicosanoids in the central nervous system. In The Eicosanoids Edited by: Curtis-Prior P. London, UK: Wiley; 210-220, 2004. [8] Carlezon surW.A Jr; Mague, S.D.; Parow, A.M.; Stoll, A.L.; Cohen, B.M.; Renshaw, P.F. Antidepressant-like effects of uridine and omega-3 fatty acids are potentiated by combined treatment in rats. Biol. Psychiatry 2005 57, 343-350. [9] Ross, B. M.; Seguin, J.; Lee E Sieswerda, L. E. Omega-3 fatty acids as treatments for mental illness: which disorder and which fatty acid? Lipids in Health and Disease 2007 6, 21. [10] Xiao, Y.F.; Sigg, D.C.; Leaf, A. The antiarrhythmic effect of n-3 polyunsaturated fatty acids: Modulation of cardiac ion channels as a potential mechanism. J. Membr. Biol., 2005 206, 141-154. [11] Wigmore, S.J.; Ross, J.A.; Falconer, J.S.; Plester, C.E.; Tisdade, M.J.; Carter, D.C.; Faron, K.C.H. Effect of PUFA on the progress of cachexia in patiens with pancreatic cancer. Nutrition, 1996 12, 27–30. [12] Beaubaire, N.A.; Simon, J.E. Production potential of Borago oficinalis L. Acta Horticulturae, 1987 208, 101-103.
176
Marie Zarevúcka and Zdeněk Wimmer
[13] Senanayake, S.P.J.N.; Shahidi, F. Enzyme-assisted acidolysis of borage (Borago officinalis L.) and evening primrose (Oenothera biennis L.) oils: Incorporation of omega-3 polyunsaturated fatty acids. J. Agric. Food Chem., 1999 47, 3105-3112. [14] Stránský, K.; Zarevúcka, M.; Wimmer, Z. Gas chromatography analysis of blackcurrant oil in relation to its stability. Food Chem. 2005 92, 569-573. [15] Traitler, H.; Wille, H.J.; Studer, A. Fractionation of Blackcurrant seed oil. J. Am. Oil Chem. Soc., 1988 65, 755-760. [16] Wille, H.J.; Traitler, H.; Lagarde, M. Preparation of stearidonic acid concentrates. Fat Sci. Technol., 1991 93, 362-368. [17] Traitler, H.; Wille, H.J. Isolation of pure fatty acids from fats and oils. Fat Sci. Technol., 1992 94, 506-511. [18] Syed Rahmatullah, M.S.K.; Shukla, V.K.S.; Mukherjee, K.D. Enrichment of γlinolenic acid from evening primrose oil and borage oil via lipase-catalyzed hydrolysis. J. Am. Oil Chem. Soc., 1994 71, 560-573. [19] Syed Rahmatullah, M.S.K.; Shukla, V.K.S.; Mukherjee, K.D. γ-Linolenic acid concentrates from borage and evening priprose oil via lipase-catalyzed esterification. J. Am. Oil Chem. Soc., 1994 71, 563-577. [20] Kawashima, H.; Sakuradani, E.; Kamada, N.; Akimoto, K.; Konishi, K.; Ogawa, J.; Shimizu, S. Production of 8,11,14,17-cis-eicosatetraenoic acid (20 : 4 omega-3) by a Delta 5 and Delta 12 desaturase-defective mutant of an arachidonic acid-producing fungus Mortierella alpina 1S-4. J. Am. Oil Chem. Soc., 1998 75, 1495-1500. [21] Shinmen, Y.; Shimizu, S.; Akimoto, D.; Kawashima, H.; Yamada, H. Production of arachidonic acid by Mortierella fungi. Selection of a potent producer and optimization of culture conditions for large-scale production. Appl. Microbiol. Biotechnol., 1989 31, 11–16. [22] Yongmanitchai, W.; Ward, O.P. Screening of algae for potential alternative sources of eicosapentaenoic acid. Phytochemistry, 1991 30, 2963–2967. [23] Nichols, B.W.; Appleby, R.S.. The distribution and biosynthesis of arachidonic acid in algae. Phytochemistry, 1969 8, 1907–1915. [24] Shimada, Y.; Sugihara, A.; Nakano, H.;Kuramoto, T.; Nagao, T.; Gemba, M.; Tominaga, Y. Purification of docosahexaenoic acid by selective esterification of fatty acids from tuna oil with Rhizopus delemar lipase. J. Am. Oil Chem. Soc., 1997 74, 97101. [25] Salunkhe, D.K.; Chavan, J. K.; Adsule, R. N.; Kadam, S. S. World oilseeds: chemistry, technology, and utilization. Ed. D. K. Salunkhe, Van Nostrand Reinhold, New York (USA) 1992. [26] Gunstone, F.D.; Ilyas-Qureshi, M. Glyceride studies. Part IV. The component glycerides of ten seed oils containing linoleic acid. J. Am. Oil. Chem. Soc., 1965 42, 961-965. [27] Gunstone, F.D.; Hamilton, R. J.; Padley, F.B.; Ilyas-Qureshi, M. Glyceride studies. V. The distribution of unsaturated acyl groups in vegetable triglycerides. J. Am. Oil. Chem. Soc., 1965 42, 965-970. [28] Biermann, U.; Friedt, W.; Lang, S.; Lühs, W.; Machmüller, G.; Metzger, J.O.; Rüsch gen. Klaas, M.; Schäfer, J.; Schneider, M.P. New syntheses with oils and fats as
Application of Lipases to Substances with Pharmacological Importance
[29] [30] [31] [32]
[33] [34]
[35] [36] [37] [38] [39] [40] [41] [42] [43] [44] [45] [46] [47]
[48]
177
renewable raw materials for the chemical industry. Angew. Chem. Int. Ed. Engl., 2000 39, 2206-2224. Hayes, D.G. Enzyme-catalyzed modification of oilseed materials to produce ecofriendly products. J. Am. Oil Chem. Soc., 2004 81, 1077-1103. Gandhi, N.N. Applications of lipase. J. Am. Oil Chem. Soc., 1997 74, 621-634. Albasi, C.; Bertrand, N.; Riba, J.P. Enzymatic hydrolysis of sunflower oil in a standardized agitated tank reactor. Bioprocess Eng., 1999 20, 77-81. Martinez, O.; Wilhelm, A.M.; Riba, J.P. Kinetic-study of an enzymatic liquid liquid reaction – the hydrolysis of tributyrin by Candida cylindracea lipase. J. Chem. Tech. Biotechnol., 1992 53, 373-378. Habulin, M.; Knez, Ž. High-pressure enzymatic hydrolysis of oil. Eur. J. Lipid Sci. Technol., 2002 104, 381-386. Malcata, F.X.; Reyes, H.R.; Garcia, H.S.; Hill Jr., C.G.; Amundson, C.H. Immobilized lipase reactions for modification of fats and oils – A review. J. Am. Oil Chem. Soc., 1990 67, 890-910. Bogel-Lukasik, R. Sustainable processes employing ionic liquids for secondary alcohols separation. Monatsh. Chem., 2007 138, 1137-1144. Dordick, J.S. Nonaqueous enzymology. Curr. Opin. Biotechnol., 1991 2, 401-407. Reverchon, E.; De Marco, I. Supercritical fluid extraction and fractionation of natural matters. J. Supercrit. Fluids, 2006 38, 146-166. Klibanov, A.M. Improving enzymes by using them in organic solvents. Nature, 2001 409, 241-246. Schoffers, E.; Golebioeski, A.; Johnson, C.R. Enantioselective synthesis through enzymatic asymmetrization. Tetrahedron, 1996 52, 3769-3826. Klibanov, A.M. Why are enzymes less active in organic solvents than in water? Trends Biotechnol., 1997 15, 97-101. Schmid, R.D.; Verger, R. Lipases: Interfacial enzymes with attractive applications. Angew. Chem. Int. Ed. Engl., 1998 37, 1608-1633. Beisson, F.; Tiss, A.; Riviere, C.; Verger, R. Methods for lipase detection and assay: a critical review. Eur. J. Lipid Sci. Technol., 2000, 133-153. Krishna, H.S.; Karanth, N.G. Lipases and lipase-catalyzed esterification reactions in nonaqueous media. Catal. Rev., 2002 44, 499-591. Patel, R.N. Microbial/enzymatic synthesis of chiral intermediates for pharmaceuticals. Enzyme Microb. Technol., 2002 31, 804-826. Patel, R.N. Biocatalytic synthesis of chiral pharmaceutical intermediates. Food Technol. Biotechnol., 2004 42, 305-325. Wong, C.-H.; Whitesides, G.M. Enzymes in Synthetic Organic Chemistry. Oxford: Pergamon; 1994. Dias, S.F.; Vilas-Boas, L.; Cabral, J.M.S.; Fonseca, M.M.R. Production of ethyl butyrate by Candida rugosa lipase immobilized in polyurethane. Biocatalysis, 1991 5, 21-34. Macrae, A.R. in: J. Tramper, H.C. van der Plas, P. Linko_Eds., Biocatalysts in Organic Synthesis, Elsevier, Amsterdam, 1985.
178
Marie Zarevúcka and Zdeněk Wimmer
[49] Muderhwa, J.M.; Pina, M.; Graille, J. Interesterification catalyzed by 1-3 regioselective lipases without solvent-valorization of palm oil and its solid fraction. Rev. Fr. Corps Gras, 1989 36, 11-19. [50] Kyotani, S.; Nakashima, T.; Izumoto, E.; Fukuda, H. Continuous interesterification of oils and fats using dried fungus immobilized in biomass support particles. J. Ferment. Bioeng., 1991 71, 286-288. [51] Mojovič, L.; Siler-Marinkovič, S.; Kukič, G.; Vunjak-Novakovič, G. Rhizopus arrhizus lipase-catalyzed interesterification of the midfraction of palm oil to a cocoa butter equivalent fat. Enzyme Microb. Technol., 1993 15, 438-443. [52] Forssell, P.; Paruvuori, P.; Linko, P.; Poutanen, K. Enzymatic transesterification of rapeseed oil and lauric acid in a continuous reactor. J. Am. Oil Chem. Soc., 1993 70, 1105-1109. [53] Ghazali, H.M.; Hamidah, S.; Che Man, Y.B. Enzymatic transesterification of palm olein with nonspecific and 1,3-specific lipases. J. Am. Oil Chem. Soc., 1995 72, 633639. [54] Soumanou, M.M.; Bornscheuer, U.T.; Menge, U.; Schmid, R.D. Synthesis of structured triglycerides from peanut oil with immobilized lipase. J. Am. Oil Chem. Soc., 1997 74, 427-433. [55] Ghosh, S.; Bhattacharyya, D.K. Utilization of high-melting palm stearin in lipasecatalyzed interesterification with liquid oils. J. Am. Oil Chem. Soc., 1997 74, 589-592. [56] Mukesh, D.; Banerji, A.A.; Newadkar, R.; Bevinakatti, H.S. Lipase catalysed transesterification of vegetable oils - A comparative study in batch and tubular reactors. Biotechnol. Lett., 1993 15, 77-82. [57] Xu, X.; Balchen, S.; Høy, C.-E.; Adler-Nissen, J. Pilot batch production of specificstructured lipids by lipase-catalyzed interesterification: Preliminary study on incorporation and acyl migration. J. Am. Oil Chem. Soc., 1998 75, 301-308. [58] Seriburi, V.; Akoh, C.C. Enzymatic transesterification of triolein and stearic acid and solid fat content of their products. J. Am. Oil Chem. Soc., 1998 75, 511-516. [59] Mu, H.; Xu, X.; Høy, C.-E. Production of specific-structured triacylglycerols by lipasecatalyzed interesterification in a laboratory-scale continuous reactor. J. Am. Oil Chem. Soc., 1998 75, 1187-1193. [60] Ferreira-Dias, S.; Duarte, C.S.; Falaschi, V.; Marques, S.R.; Gusmao, J.H.; da Fonseca, M.M.R. in: A. Ballesteros, F.J. Plou, J.L. Iborra, P. Halling-Eds., Stability and Stabilization of Biocatalysts, Elsevier, 1998. [61] Fomuso, L.B. ; Akoh, C.C. Enzymatic modification of triolein: Incorporation of caproic and butyric acids to produce reduced-calorie structured lipids. J. Am. Oil Chem. Soc., 1997 74, 269-272. [62] Xu, X. ; Skands, A.R.H. ; Høy, C.-E. ; Mu, H.; Balchen, S.; Adler-Nissen, J. Production of specific-structured lipids by enzymatic interesterification: Elucidation of acyl migration by response surface design. J. Am. Oil Chem. Soc., 1998 75, 1179-1186. [63] Miura, S.; Ogawa, A.; Konishi, H. A rapid method for enzymatic synthesis and purification of the structured triacylglycerol, 1,3-dilauroyl-2-oleoyl-glycerol. J. Am. Oil Chem. Soc., 1999 76, 927-931.
Application of Lipases to Substances with Pharmacological Importance
179
[64] Adachi, S.; Okumura, K.; Ota, Y.; Mankura, M. Acidolysis of sardine oil by lipase to concentrate eicosapentaenoic and docosahexanoic acids in glycerides. J. Ferment. Bioeng., 1993 75, 259-264. [65] Yamane, T.; Suzuki, T.; Hoshino, T. Increasing n-3 polyunsaturated fatty acid content of fish oil by temperature control of lipase-catalyzed acidolysis J. Am. Oil Chem. Soc., 1993 70, 1285-1287. [66] Zuyi, L.; Ward, O.P. Enzyme catalysed production of vegetable oils containing omega3 polyunsaturated fatty acid. Biotechnol. Lett., 1993 15, 185-188. [67] Huang, K.-H.; Akoh, C.C. Lipase-catalyzed incorporation of n−3 polyunsaturated fatty acids into vegetable oils. J. Am. Oil Chem. Soc., 1994 71, 1277-1288. [68] Lee, K.-T.; Akoh, C.C. Immobilized lipase-catalyzed production of structured lipids with eicosapentaenoic acid at specific positions. J. Am. Oil Chem. Soc., 1996 73, 611615. [69] Senanayake, S.P.J.N.; Shahidi, F. Enzymatic incorporation of docosahexaenoic acid into borage oil. J. Am. Oil Chem. Soc., 1999 76, 1009-1015. [70] Jennings, B.H.; Akoh, C. Enzymatic modification of triacylglycerols of high eicosapentaenoic and docosahexaenoic acids content to produce structured lipids. J. Am. Oil Chem. Soc., 1999 76, 1133-1137. [71] Haraldsson, G.; Thorarensen, A. Preparation of phospholipids highly enriched with n-3 polyunsaturated fatty acids by lipase. J. Am. Oil Chem. Soc., 1999 76, 1143-1149. [72] Muderhwa, J.M.; Pina, M.; Montet, D.; Feuillard, P.; J. Graille, 1-3 Regioselective enzymatic interesterification in a melted medium and a continuous reactor – valorization of palm oil. Oleagineux, 1989 44, 36-43. [73] Zainal, Z.; Yusoff, M.S.A. Enzymatic interesterification of palm stearin and palm kernel olein. J. Am. Oil Chem. Soc., 1999 76, 1003-1008. [74] Bergstrom, B.; Brochmann, H. L. Lipases, 1st ed. Elsevier, Amsterdam, New York (1984). [75] Quinlan, P.; Moore, S. Modification of triglycerides by lipases: process technology and its application to the production of nutritionally improved fats. INFORM, 1993 4, 580585. [76] Brady, L.; Brzozowski, A.M.; Derewenda, Z. S.; Dodson, E.; Dodson, G.; Tolley, S.; Turkenburg, J. P.; Christiansen, L.; Huge-Jensen, B.; Norskov, L.; Thim, L.; Menge, U. A serine protease triad forms the catalytic centre of a triacylglycerol lipase. Nature, 1990 343, 767-770. [77] Klibanov, A.M. Why are enzymes less active in organic solvents than in water? Trends Biotechnol. 1997 15, 97-101. [78] Pleiss , J.; Fischer, M.; Peiker, M.; Thiele, C.; Schmid, R.D. Lipase engineering database. Understanding and exploiting sequence–structure–function relationships. J. Mol. Catal. B Enzymatic, 2000 10, 491–508. [79] Balcao V.M.; Paiva, A.L.; Malcata, F.X. Review. Bioreactors with immobilized lipases: state of the art. Enzyme Microbial. Technol., 1996 18, 392–416. [80] Deng, H.T.; Xu, Z.K.; Huang, X.J.; Wu, J.; Seta, P. Adsorption and activity of Candida rugosa lipase on polypropylene hollow fiber membrane modified with phospholipid analogous polymers. Langmuir, 2004 20, 10168–10173.
180
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[81] Homquist, M.; Norin, M.; Hult, K. The role of arginines in stabilizing the active openlid conformation of Rhizomucor meihei lipase. Lipids, 1993 28, 721–726. [82] Villeneuve, P.; Muderwha, J.; Graille, J.; Haas, M. Customizing lipases for biocatalysis: a survey of chemical, physical and molecular biological approaches. J. Mol. Catal. B Enzymatic, 2000 9, 113–148. [83] Holmquist, M.; Martinelle, M.; Berglund, P.; Clausen, I.G.; Patkar, S.; Svendsen, A.; Hult, K. Lipases from Rhizomucor miehei and Thermomyces lanuginosa: modification of the lid covering the active site alters enantioselectivity. J. Protein Chem., 1993 12, 749–757. [84] Gutierrez-Ayesta, C.; Carelli, A.A.; Ferreira, M.L. Relation between lipase structures and their catalytic ability to hydrolyse triglycerides and phospholipids. Enzyme Microb. Technol., 2007 41, 35–43. [85] Pleiss, J.; Fisher, M.; Schmid, R.D. Anatomy of lipase binding sites: the scissile fatty acid binding site. Chem. Phys. Lipids, 1998 93, 67–80. [86] Bottino, N.R.; Vandenburg, G.A.; Reiser, R. Resistance of certain long chain polyunsaturated fatty acids of marine oils to pancreatic lipase hydrolysis. Lipids, 1967 2, 489-493. [87] Jensen, R.G.; Gordon, D.T. Specificity of Geotrichum candidum lipase with respect to double bond position in triglycerides containing cis-octadecenoic acids. Lipids, 1972 7, 738-741. [88] Lopez-Martinez, J.C.; Campra-Madrid, P.; Ramirez-Fajardo, A.; Esteban-Cerdan, L.; Guil-Guerrero, J.L. Screening of lipases for enzymatic concentration of gammalinolenic acid (GLA) from seed oils. J. Food Lipids, 2006 13, 362-374. [89] Hoshino, T.; Yamane, T.; Shimizu, S. Selective hydrolysis of fish oil by lipase to concentrate n-3 polyunsaturated fatty acid. Agric. Biol. Chem., 1990 54, 1459-1467. [90] Tanaka, Y.; Hirano, J.; Funada, T. Concentration of docosahexaenoic acid glyceride by hydrolysis of fish oil with Candida cylindracea lipase. J. Am. Oil Chem. Soc., 1992 69, 1210-1214. [91] Basheer, S.; Plat, D. Enzymatic modification of sterols using sterol-specific lipase. WO patent 2001-075083; 2001. [92] Weber, N.; Weikamp, P.; Mukherjee, K.D. Cholesterol-lowering food additives: Lipase-catalyzed preparation of phytosterol and phytostanol esters. Food Res. Intl., 2002 35, 177-181. [93] Bertinotti, A.; Carrea, G.; Ottolina, G.; Riva, S. Regioselective esterification of polyhydroxylated steroids by Candida antarctica lipase B. Tetrahedron, 1994 50, 13165-13172. [94] Norinobu, S.; Senoo, N.; Kaneko, S.; Sato, F; Mankura, M. Supercritical preparation of sterol fatty esters with enzyme. Japanese patent 2002-233396; 2002. [95] Stamatis, H.; Sereti, V.; Kolisis, F.N. Studies on the enzymatic synthesis of lipophilic derivatives of natural antioxidants. J. Am. Oil Chem. Soc., 1999 76, 1505-1510. [96] Jessop, P.G.; Ikariya, T.; Noyori, R. Homogenous catalysis in supercritical fluids. Chem. Rev., 1999 99, 475-494. [97] Brennecke, J.F.; Chateauneuf, J.E. Homogenous organic reactions as mechanistic probes in supercritical fluids. Chem. Rev., 1999 99, 433-452.
Application of Lipases to Substances with Pharmacological Importance
181
[98] Baiker, A. Supercritical fluids in heterogenous catalysis. Chem. Rev., 1999 99, 453474. [99] Darr, J.A.; Poliakoff, M. New directions in inorganic and metal-organic coordination chemistry in supercritical fluids. Chem. Rev., 1999 99, 495-542. [100] Kiran, E.; Debenedetti, P.G.; Peters, C.J. Supercritical Fluids Fundamentals and Applications; Dordrecht: Kluwer Academic; 2000. [101] Leitner, W. Supercritical carbon dioxide as a green reaction medium for catalysis. Acc. Chem. Res., 2002 35, 746-756. [102] Matsuda, T.; Watanabe, K.; Harada, T.; Nakamura, K. Enzymatic reactions in supercritical CO2: carboxylation, asymmetric reduction and esterification. Catal. Today, 2004 96, 103-111. [103] Kragl, U.; Eckstein, M.; Kraftzik, N. Biocatalytic Reactions in Ionic Liquids. In: Wasserscheid, P.; Welton, T., editors. Ionic Liquids in Synthesis. Weinheim: WileyVCH; 2003; 336-347. [104] Seddon, K.R. Ionic liquids for clean technology. J. Chem. Technol. Biotechnol., 1997 68, 351-356. [105] Welton, T. Room-temperature ionic liquids. Solvents for synthesis and catalysis. Chem. Rev., 1999 99, 2071-2084. [106] Yang, Z.; Pan, W. Ionic liquids: green solvents for nonaqueous biocatalysis. Enzyme Microb. Technol., 2005 37, 19-28. [107] Chowdhury, S.; Mohan, R.S.; Scott, J.L. Reactivity in ionic liquids. Tetrahedron, 2007 63, 2363-2389. [108] Freemantle, M. Ionic liquids in organic synthesis. Chem. Eng. News, 2004 82, 44-49. [109] Roberts, N.J.; Seago, A.; Carey, J.S.; Freer, R.; Preston, C.; Lye, G.J. Lipase catalysed resolution of the Lotrafiban intermediate 2,3,4,5-tetrahydro-4-methyl-3-oxo-1H-1,4benzodiazepine-2-acetic acid methyl ester in ionic liquids: comparison to the industrial t-butanol process. Green Chem., 2004 6, 475-482. [110] Lau, R.M.; Sorgedrager, M.J.; Carrea, G.; van Rantwijk, F.; Secundo, F.; Sheldon, R.A. Dissolution of Candida antarctica lipase B in ionic liquids: effects on structure and activity. Green Chem., 2004 6, 483-487. [111] Oliveira, J.V.; Oliveira, D. Kinetics of the enzymatic alcoholysis of palm kernel oil in supercritical CO2. Ind. Eng. Chem. Res., 2000 39, 4450-4454. [112] Kamat, S.V.; Beckman, E.J.; Russell, A.J.: Enzyme-activity in supercritical fluids. Crit. Rev. Biotechnol., 1995 15, 41-71. [113] Connstein, W.; Hoyer, E.; Wartenberg, H. The enzymic decomposition of fat. Chem. Ber., 1902 34, 3988-4006. [114] Baumann, H.; Bühler, M.; Fochem, H.; Hirsinger, F.; Zoebelein, H.; Falbe, J. Natural Fats and Oils - Renewable Raw Materials for the Chemical Industry. Angew. Chem. Int. Ed. Engl., 1988 27, 41-62. [115] Moquin, P.H.L.; Temelli, F. Production of monoolein from oleic acid and glycerol in supercritical carbon dioxide media: a kinetic approach. J. Supercrit. Fluids, 2008 44, 40-47. [116] Promožič, M.; Habulin, M.; Knez, Ž. Parameter optimization for enzymic hydrolysis of sunflower oil in high-pressure reactors. J. Am. Oil Chem. Soc., 2003 80, 643-646.
182
Marie Zarevúcka and Zdeněk Wimmer
[117] Promožič, M.; Habulin, M.; Knez, Ž. Thermodynamic properties of the enzymatic hydrolysis of sunflower oil in high-pressure reactors. J. Am. Oil Chem. Soc., 2003 80, 785-788. [118] Promožič, M.; Habulin, M.; Knez, Ž. Modeling of kinetics for the enzymatic hydrolysis of sunflower oil in a high-pressure reactors. J. Am. Oil Chem. Soc., 2005 82, 543-547. [119] Zarevúcka, M.; Vacek, M.; Wimmer, Z.; Stránský, K.; Koutek, B.; Demnerová, K. Enzymic transformations of blackcurrant oil: enrichment with γ-linolenic acid and αlinolenic acid. Chem. Listy, 2003 97, 206-213. [120] Sovová, H.; Zarevúcka, M. Lipase-catalysed hydrolysis of blackcurrant oil in supercritical carbon dioxide. Chem. Eng. Sci., 2003 58, 2339-2350. [121] Bártlová, M.; Bernášek, P.; Sýkora, J.; Sovová, H. HPLC in reversed phase mode: tool for investigation of kinetics of blackcurrant seed oil lipolysis in supercritical carbon dioxide. J. Chromatogr. B, 2006 839, 80-84. [122] Sýkora, J.; Bernášek, P.; Zarevúcka, M.; Kurfürst, M.; Sovová, H.; Schraml, J. Highperformance liquid chromatography with nuclear magnetic resonance detection - A method for quantification of α- and γ-linolenic acids in their mixtures with free fatty acids. J. Chromatogr. A, 2007 1139, 152-155. [123] Kosugi, Y.; Azuma, N.; Synthesis of triacylglycerol from polyunsaturated fatty acid by immobilized lipase. J. Am. Oil Chem. Soc., 1994 71, 1397–1403. [124] Cerdán, L.E.; Medina, A.R.; Giménez Giménez, A.; Ibánez González, M.J.; Molina Grima, E. Synthesis of PUFA enriched triglycerides by lipase catalyzed esterification. J. Am. Oil Chem. Soc., 1998 75, 1329–1337. [125] Medina, A.R.; Cerdán, L.E.; Giménez Giménez, A.; Camacho Páez, B.; Ibánez González, M.J.; Molina Grima, E. Lipase-catalyzed esterification of glycerol and polyunsaturated fatty acids from fish and microalgae oils. J. Biotechnol. 1999 70, 379– 391, [126] Akoh, C.C.; Moussata, C.O. Lipase-catalyzed modification of borage oil: Incorporation of capric and eicosapentaenoic acids to form structured lipids. J. Am. Oil Chem. Soc., 1998 75, 697-701. [127] McKenna, M. C.; Hubbard, V. S.; Aeri, J. G. Linoleic acid absorption for lipid supplements in patients with cystic fibrosis with pancreatic insufficiency and in control subjects. J. Pediatr. Gastroenterol. Nutr., 1985 4, 45–48. [128] Kawashima, A.; Shimada, Y.; Nagao, T.; Ohara, A.; Matsuhisa, T.; Sugihara, A.; Tominaga, Y. Production of structured TAG rich in 1,3- dicapryloyl-2-c-linolenoyl glycerol from borage oil. J. Am. Oil Chem. Soc., 2002 79, 871–877. [129] Kawashima, A.; Shimada, Y.; Yamamoto, M.; Sugihara, A.; Nagao, T.; Komemushi, S.; Tominaga, Y. Enzymatic synthesis of high-purity structured lipids with caprylic acid at 1,3-positions and polyunsaturated fatty acid at 2-position. J. Am. Oil Chem. Soc., 2001 78, 611–616. [130] Senanayake, S.P.J.N.; Shahidi, F. Chemical and stability of structured lipids from borage (Borage officinalis L.) and evening primrose (Oenothera biennis L.) oils. J. Food Sci., 2002 67, 2038–2045.
Application of Lipases to Substances with Pharmacological Importance
183
[131] Shimada, Y.; Suenaga, M.; Sugihara, A.; Seiichi, S.; Tominaga, Y. Continuous production of structured lipids containing clinolenic and caprylic acids by immobilized Rhizopus delemar lipase. J. Am. Oil Chem. Soc., 1996 76, 189–193. [132] Hamam, F.; Shahidi, F. Synthesis of structured lipids via acidolysis of docosahexaenoic acid single cell oil (DHASCO) with capric acid. J. Agric. Food Chem., 2004 52, 2900–2906. [133] Hamam, F.; Shahidi, F. Enzymatic acidolysis of arachidonic acid single cell oil (ARASCO) with capric acid. J. Am. Oil Chem. Soc., 2004 81, 887–892. [134] Hamam, F.; Shahidi, F. Enzymatic incorporation of capric acid into a single cell oil rich in docosahexaenoic acid (DHA) and docosapentaenoic acid (DPA). Food Chem., 2005 91, 583–591. [135] Iwasaki, Y.; Han, J. J.; Narita, M.; Rosu, R.; Yamane, T. Enzymatic synthesis of structured lipids from single cell oil of high docosahexaenoic acid content. J. Am. Oil Chem. Soc., 1999 76, 563–569. [136] Yang, T.; Xu, X.; Li, L. Comparison of linoleic and conjugated linoleic acids in enzymatic acidolysis of tristearin. J. Food Lipids, 2001 8, 149–161. [137] Ronne, T. H.; Pederson, L. S.; & Xu, X. Triglyceride selectivity of immobilized thermomyces lanuginose lipase in interesterification. J. Am. Oil Chem. Soc., 2005 82, 737–743. [138] Tsuzuki, W. Acidolysis between triolein and short-chain fatty acids by lipase in organic solvents. Biosci. Biotechnol., Biochem., 2005 69, 1256–1261. [139] Paez, B. C.; Medina, A. R.; Rubio, F. C.; Cerdan, L. E.; Grima, E. M. Kinetics of lipase-catalyzed interesterification of triolein and caprylic acid to produce structured lipids. J. Chem. Technol. Biotechnol., 2003 78, 461–470. [140] Sellappan, S.; Akoh, C. C. Synthesis of structured lipids by transesterification of trilinolein catalyzed by Lipozyme IM60. J. Agric. Food Chem., 2001 49, 1–9. [141] Fomus, L.B.; Akoh, C.C. Structured lipids: Lipase-catalyzed interesterification of tricaproin and trilinolein. J. Am. Oil Chem. Soc., 1998 75, 405–410. [142] Hamam, F.; Shahidi F. Incorporation of selected long-chain fatty acids into trilinolein and trilinolenin. Food Chem., 2008 106, 33–39. [143] Hamam, F.; Shahidi, F. Enzymatic incorporation of selected long-chain fatty acids into triolein. J. Am. Oil Chem. Soc., 2007 84, 533-541. [144] Hamam, F.; Shahidi, F. Acidolysis of tristearin with selected long chain fatty acids. J. Agric. Food Chem., 2007 55, 1955–1960. [145] Lee, K.; Akoh C. Characterization of enzymatically synthesized structured lipids containing eicosapentaenoic, docosahexaenoic, and caprylic acids. J. Am. Oil Chem. Soc., 1998 75, 495-499. [146] McNeill, G. P.; Ackman, R. G.; Moore S. R. Lipase-catalyzed enrichment of long chain polyunsaturated fatty acids. J. Am. Oil Chem. Soc., 1996 73, 1403-1414. [147] Schmid, U.; Bornscheuer, U. T.; Soumanou, M. M.; Mc- Neill, G. P.; Schmid, R. D. Optimization of the reaction conditions in the lipase-catalyzed synthesis of structured triglycerides. J. Am. Oil Chem. Soc., 1998 75, 1527-1531. [148] Soumanou, M.M.; Bornscheuer, U.T.; Schmid R.D. Two step enzymatic reaction for synthesis of pure structured triacylglycerides. J. Am. Oil Chem. Soc., 1998 75, 703 710.
184
Marie Zarevúcka and Zdeněk Wimmer
[149] Tsujisaka, Y.; Okumura, S.; Iwai, M. Glyceride synthesis by four kinds of microbial lipase. Biochim. Biophys. Acta, 1977 489, 415-422. [150] Okumura, S.; Iwai, M.; Tsujisaka, Y. Synthesis of various kinds of esters by four microbial lipases. Biochim. Biophys. Acta, 1979 575, 156-165. [151] Liu, K.-J.; Chang, H.-M.; Liu, K.-M. Enzymatic synthesis of cocoa butter analog through interesterification of lard and tristearin in supercritical carbon dioxide by lipase. Food Chem., 2007 100, 1303-1311. [152] Jessop, P.; Leitner, W. Chemical synthesis using supercritical fluids. Weinheim: Wiley-VCH; 1999. [153] King, J.W.; Snyder, J.M.; Frykman, H.; Neese, A. Sterol ester production using lipasecatalyzed reactions in supercritical carbon dioxide. Eur. Food Res. Technol., 2001 212, 566-569. [154] Gunnlaugsdottir, H.; Järemo, M.; Sivik, B. Process parameters influencing ethanolysis of cod liver oil in supercritical carbon dioxide. J. Supercrit. Fluids, 1998 12, 85-93. [155] Overmeyer, A.; Schrader-Lippelt, S.; Kasche, V.; Brunner, G. Lipase-catalysed kinetic resolution of racemates at temperatures from 40 °C to 160 °C in supercritical CO2. Biotechnol. Lett., 1999 21, 65-69. [156] Habulin, M.; Krmelj, V.; Knez, Ž. Synthesis of oleic acid esters catalyzed by immobilized lipase. J. Agric. Food Chem., 1996 44, 338-342. [157] Knez Ž.; Habulin, M. Compressed gases as alternative enzymatic-reaction solvents: a short review. J. Supercrit. Fluids, 2002 23, 29-42. [158] Leitner, W. Die “bessere Lösung”? Chemische Synthese in überkritischem Kohlendioxid. Chem. Unserer Zeit, 2003 37, 32-38. [159] King, J.; Jackson, M.; Temelli, F. (1995) Development of industrially-useful synthetic processes in supercritical carbon dioxide. In: Kikic, I.; Alessi, P., Editors. I Fluidi Supercritici e le Lorno Applicazioni. Trieste; 1995; 19-26. [160] Bornscheuer, U.; Kazlauskas, R. Hydrolases in organic synthesis: regio- and stereoselective biotransformations. Weinheim: Wiley-VCH; 1999. [161] Snyder, J.; King, J.; Jackson, M. Fat content for nutritional labeling by supercritical fluid extraction and an on-line lipase catalyzed reaction. J. Chromatogr. A, 1996 750, 201-207. [162] Anderson, M.; King, J.; Blomberg, L. Synthesis of fatty alcohol mixtures from oleochemicals in supercritical fluids. Green Chem., 2000 2, 230-234. [163] Nakaya, H.; Nakamura, K.; Miyawaki, O. Lipase-catalyzed esterification of stearic acid with ethanol, and hydrolysis of ethyl stearate, near the critical point in supercritical carbon dioxide. J. Am. Oil Chem. Soc., 2002 79, 23-27. [164] Ikushima, Y.; Saito, N.; Arai, M.; Blanch, H.W. Activation of lipase triggered by interactions with supercritical carbon dioxide in the near-critical region. J. Phys. Chem., 1995 99, 8941-8944. [165] Srivastava, S.; Madras, G.; Modak, J. Esterification of myristic acid in supercritical carbon dioxide. J. Supercrit. Fluids, 2003 27, 55-64. [166] Gunnlaugsdottir, H.; Wannerberger, K.; Sivik, B. Alcoholysis and glyceride synthesis with immobilized lipase on controlled-pore glass of varying hydrophobicity in supercritical carbon dioxide. Enzyme Microb. Technol., 1998 22, 360-367.
Application of Lipases to Substances with Pharmacological Importance
185
[167] Laudani, C.G.; Habulin, M.; Primožič, M.; Knez, Ž.; Della Porta, G.; Reverchon, E. Optimisation of n-octyl oleate enzymatic synthesis over Rhizomucor miehei lipase. Bioprocess Biosyst. Eng., 2006 29, 119-127. [168] Laudani, C.G.; Habulin, M.; Knez, Ž.; Della Porta, G.; Reverchon, E. Immobilized lipase-mediated long-chain fatty acid esterification in dense carbon dioxide: benchscale packed-bed reactor study. J. Supercrit. Fluids, 2007 41, 74-81. [169] Pomier, E.; Galy, J.; Paolucci-Jeanjean, D.; Pina, M.; Sarrade, S.; Rios, G.M. A new reactor design combining enzyme, membrane and SC CO2: application to castor oil modification. J. Membrane Sci., 2005 249, 127-132. [170] Liu, K.-J.; Liu, K.-M.; Chang, H.-M. Biocatalytic synthesis of palmitoyl vanillylamide in supercritical carbon dioxide through amidation of vanillylamide hydrochloride and palmitic anhydride by lipase. Food Chem., 2007 102, 1020-1026. [171] Hakoda, M.; Shiragami, N.; Enomoto, A.; Nakamura, K. Effect of moisture on enzymatic reaction in supercritical carbon dioxide. Bioprocess Biosyst. Eng., 2002 24, 355-361. [172] Eisenmenger, M.; Dunford, N.T.; Eller, F.; Tailor, S.; Martinez, J. Pilot-scale supercritical carbon dioxide extraction and fractionation of wheat germ oil. J. Am. Oil Chem. Soc., 2006 83, 863-868. [173] Lau, H.L.N.; Choo, Y.M.; Ma, A.N.; Chuah, C.H. Quality of residual oil from palmpressed mesocarp fiber (Elaeis guineensis) using supercritical CO2 with and without ethanol. J. Am. Oil Chem. Soc., 2006 83, 893-898. [174] Üstündag, Ö.G.; Temelli, F. Column fractionation of canola oil deodorizer distillate using supercritical carbon dioxide. J. Am. Oil Chem. Soc., 2007 84, 953-961. [175] Nagesha, G.K.; Manohar, B.; Sankar, K.U. Enzymatic esterification of free fatty acids of hydrolyzed soy deodorizer distillate in supercritical carbon dioxide. J. Supercrit. Fluids, 2004 32, 137-145. [176] Frykman, H.B.; Snyder, J.M.; King, J.W. Screening catalytic lipase activities with an analytical supercritical fluid extractor. J. Am. Oil Chem. Soc., 1998 75, 517-520. [177] Hernández, F.J..; de los Ríos, A.P.; Gómez, D.; Rubio, M.; Víllora, G. A new recirculating enzymatic membrane reactor for ester synthesis in ionic liquid/supercritical carbon dioxide biphasic systems. Appl. Catal. B: Environ., 2006 67, 121-126. [178] Gumí, T.; Paolucci-Jeanjean, M.; Belleville, M.-P.; Rios, G.M. Enzymatic membrane reactor involving a hybrid membrane in supercritical carbon dioxide. J. Membrane Sci., 2007 297, 98-103. [179] Wang, L.; Weller, C.L. Recent advances in extraction of nutraceuticals from plants. Trends Food Sci. Technol., 2006 17, 300-312. [180] Ghanem, A. Trends in lipase-catalyzed asymmetric access to enantiomerically pure/enriched compounds. Tetrahedron, 2007 63, 1721-1754. [181] Jurček, O.; Wimmerová, M.; Wimmer, Z. Selected Chiral Alcohols: Enzymic Resolution and Reduction of Convenient Substrates. Coord. Chem. Rev., 2008, in press.
In: Biocatalysis Research Progress Editors: F. H. Romano, A. Russo
ISBN: 978-1-60456-619-2 © 2008 Nova Science Publishers, Inc.
Chapter V
The Evolution of Directed Evolution Birthe Borup, Lynne Gilson, Richard Fox and Thomas Daussmann Codexis Laboratories Singapore Pte Ltd
Abstract The recent increasing usage of biocatalysts as viable alternatives to conventional chemical reactions is attributable to improved technology in tailoring enzymes found in nature to the needs of the industry. Arguably the most successful technology in this field has been directed evolution. Here we present the various stages in the evolution of directed evolution itself. The initial approach employed in directed evolution was successive rounds of random mutagenesis and screening, from which the top performer was chosen as parent for the next round of mutagenesis. Although effective, this method is not very efficient, as all beneficial mutations not found in the top performer are lost in the next round of evolution. With the advent of DNA recombination methods, mutations found in lower performing enzymes could be included in the next round. However, there was still useful diversity left unrecognized and unutilized, since only the top performing enzymes would be used for recombination. As statistical tools based on sequence-activity relationships were developed to help identify good mutations for recombination, beneficial diversity from even poorly performing enzymes could be moved to the next round of evolution. Presently, the drive to increase knowledge of sequence-activity relationships for enzyme families is enabling collections of variants to be synthesized that efficiently explore the chemical space of potential substrates for those enzymes. This will continue to lead to even faster evolution cycles, as a more relevant portion of the sequence space to be explored has been designed into the first library (first set of enzymes to be screened).
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Introduction Biocatalysts are increasingly being used in a wide variety of industries, from bioindustrial (e.g. pulp and paper, renewable and clean energies) to life sciences (pharma, food, feed, agro). The exquisite regio- and chemo- and enatioselectivity of enzymes make them especially advantageous for the synthesis of chiral compounds. They are also becoming of increasing interest in designing more environmentally benign processes and for bioremediation [1-3]. However there is usually a wide gap between what an enzyme found in nature is capable of catalyzing, and what is required for a cost effective industrial process, since substrate specificity, substrate or product inhibition, enantioselectivity, rate of catalysis, pH optimum, and stability at elevated temperatures and in organic solvents, will be very specific for any desired reaction. Historically, the most effective technique used to overcome these obstacles is the directed evolution of an enzyme, during which the enzyme is tailored for a specific chemical process through iterative rounds of variant creation and screening for desirable properties.
Random Mutagenesis In a pioneering example of directed molecular evolution from the 1960s, Spiegleman and coworkers [4] were able to directly evolve an RNA molecule to increase its speed of replication by applying a simple selection pressure for molecules which duplicated faster in vitro and where random mutations were the only source of genetic variability. Though examples of directed enzyme evolution can be found as early as the 1970s using classical strain improvement techniques involving growth under mutagenic conditions [5, 6], more targeted methods similar to that done earlier with RNA molecules did not achieve widespread use until the advent of gene level random mutagenesis became available [7, 8]. Generation of these types of libraries does not require any additional information about the enzyme, such as structure, active site, or enzyme mechanism. Sequencing of the best variant at the end of each round of evolution is not necessary, as only the top performer is subjected to another round of random mutagenesis. A variety of random mutagenesis methods have been developed over the years, such as chemical mutagens, reverse transcriptase, error-prone rolling circle amplification, error prone PCR (epPCR), and mutazyme. [9]. Currently, the most popular form of random mutagenesis is error prone PCR (epPCR), in which mistakes are introduced into a gene during DNA polymerase-catalyzed amplification cycles. After one round of random mutagenesis and screening, the top performer is selected and subjected to another run of epPCR to add additional mutations. This method has proven successful in evolving a variety of enzymes for a variety of traits, such as increase in product yield of a toluene dioxygenase [10], enantioselectivity of an epoxide hydrolase [11] and activity in organic solvents of subtilisin [12].
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Site-direct Mutations and Randomizations Through a variety of techniques, it is possible to identify amino acid residues that could affect catalysis or stability of a biocatalyst. Sequencing of improved variants from a round of random mutagenesis leads to the identification of mutations that have an effect on catalysis or stability. Alignment of homologous genes can also help with the identification of residues. Computational docking studies of a substrate, product, and transition state into the active site of an enzyme, in conjunction with knowledge of the enzymes mechanism, are a powerful method to identify amino acid residues that are potentially important in catalysis. In the past 15 years, the number of crystal structures available through the Protein Data Bank has increased exponentially. The number of available structures, faster computers and better techniques have made homology modeling of enzymes with as low as 30% sequence identity with a known structure possible [13]. Once the residues have been identified, these positions can be mutated to either specific amino acids [8], a small library of changes, or randomized to all 20 possible amino acids, via degenerate or pooled oligonucleotides [14, 15]. After screening of the new variants, the top performer is selected, and subjected to a new round of targeted randomization. This method has been effectively applied on many enzymes. For example, it has been used to increase the activity of transketolase using structural proximity and phylogenetic variation to determine positions of interest [16], iterative saturation mutagenesis was used to increase the thermostability of lipase, using crystal structures to determine areas of the genes with the highest B-factor [17] (i.e. the area of a crystal structure containing the highest uncertainty of the atomic position, either due to flexibility in the protein or error in the model), and structure guided saturation and site directed mutagenesis was used to change the N-acetylneuraminic acid lyase (NAL) from a non-selective enzyme to two catalysts with complementary stereoselectivity [18].
DNA Recombination The main drawback in using random mutagenesis and site-directed randomizations as the only tool for directed evolution, is that after each round of screening, only the top performer is chosen as the backbone for the next round. Good mutations in enzymes that were not in the top performer get lost and have to be rediscovered [19]. DNA recombination techniques enable the recombination of beneficial mutations from the top tier of enzymes, resulting in larger increases in desired enzyme properties [20]. The larger the variety of beneficial mutations that are found in the top performers, the easier it becomes to increase desired enzyme properties. This shifts the focus away from the top performers and towards the collection of mutations found in the population (also referred to as the diversity) [21]. A variety of DNA recombination methods have been developed over the years, such as DNA shuffling, exon-shuffling, RACHITT, CLERY, SHIPREC, ITCHY, THIO-ITCHY, and SCRATCHY [21, 22]. If no additional mutations are added after each round of evolution, the diversity pool decreases, since only the diversity in the top performers makes it to the next round. This makes it increasingly difficult to isolate enzymes with increases in desired
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enzyme properties. Therefore, a combination of site-directed mutagenesis, random mutagenesis, and DNA shuffling becomes the best strategy for directed evolution. DNA recombination techniques have been successfully applied to increase more than one trait in an enzyme, such as increased activity of lignin peroxidase on 2,4-dichlorophenol degradation as well as increased stability towards peroxide [23], and increased activity of deacetoxycephalosporin C synthase (expandase) on penicillin G with no substrate inhibition [24]. The technique has also been applied to achieve remarkable activity enhancements, such as the evolution of glyphosate acetyl transferase (GAT) to four orders of magnitude increase in activity in 11 rounds of evolution.
Shuffling of Mutations With the advent of cheap sequencing (the cost of sequencing dropped from $1 per base in the 1980’s to mere cents per base today), more clones than just the top hits could be sequenced. This resulted in a shift in focus from diversity pools to specific mutations. This opened the field to analyze the opposing pull of the diversity pool vs. individual mutations. That is, it became apparent that including as many hits as possible in the shuffling reaction (to keep the diversity pool high) leads to the diluting out of individual mutations (since the probability of finding any specific mutation in the progeny is the number of parents it is found in divided by the total number of parents in the shuffling reaction) [25]. This led to the development of semi-synthetic shuffling in which oligos carrying the targeted mutations are added to the shuffling reaction instead of the entire gene on which the mutation was found. This method enables maximizing the amount of diversity that is moved to the next round but avoids the dilution effect, since the probability of finding any specific mutation in the progeny is proportional to the oligo concentration [26, 27]. With small data sets it is possible to discern which mutations are beneficial for a given trait, by manual manipulation of the data. However, as more mutations are evaluated in a given library, and each library is screened for a variety of traits, the data set becomes too large to analyze by hand. Statistical models, such as ProSAR, were developed to help analyze this data [28, 29], and were successfully applied to the evolution of a haloydrin dehalogenase [30].
The Future of Directed Evolution Directed evolution has proved to be a powerful tool in generating biocatalysts that are tailor-made for industrial applications. Directed evolution itself has evolved from blindly mutating genes and screening for improved activity towards a comprehensive understanding of the effect of individual mutations in a given protein. The future in evolution will rely on using this information gained on individual enzyme classes to develop panels of enzymes that explore the sequence space efficiently. Testing of these enzyme panels not only helps in identifying a starting point for evolution, but through statistical analysis of the data, can also predict a subset of enzymes that will perform better than any of the enzymes on the panel.
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This eliminates time lost in generating and screening enzyme libraries and brings biocatalysts closer to the market faster.
References [1]
[2]
[3] [4]
[5]
[6]
[7]
[8] [9]
[10]
[11]
[12]
Alcalde, M., Ferrer, M., Plou, F. J., and Ballesteros, A. (2006) Environmental biocatalysis: from remediation with enzymes to novel green processes, Trends Biotechnol 24, 281-287. Grate, J. (2006) Directed Evolution of Three Biocatalysts to Produce the Key Chiral Building Block for Atorvastatin, the Active Ingredient in Lipitor®, in 2006 Presidential Green Chemisty Challenge Award: Greener Reaction Conditions Award, United States Environmental Protection Agency, Washington, D.C. Pollard, D. J., and Woodley, J. M. (2007) Biocatalysis for pharmaceutical intermediates: the future is now, Trends Biotechnol 25, 66-73. Mills, D. R., Peterson, R. L., and Spiegelman, S. (1967) An extracellular Darwinian experiment with a self-duplicating nucleic acid molecule, Proc Natl Acad Sci U S A 58, 217-224. Pabst, M. J., Kuhn, J. C., and Somerville, R. L. (1973) Feedback regulation in the anthranilate aggregate from wild type and mutant strains of Escherichia coli, J Biol Chem 248, 901-914. Scazzocchio, C., and Sealy-Lewis, H. M. (1978) A mutation in the xanthine dehydrogenase (purine hydroxylase I) of Aspergillus nidulans resulting in altered specificity. Implications for the geometry of the active site, Eur J Biochem 91, 99109. Leung, D. W., Chen, E., and Goeddel, D. V. (1989) A method for random mutagenesis of a defined DNA segment using a modified polymerase chain reaction, Technique 1, 11-15. Botstein, D., and Shortle, D. (1985) Strategies and applications of in vitro mutagenesis, Science 229, 1193-1201. Wong, T. S., Roccatano, D., Zacharias, M., and Schwaneberg, U. (2006) A statistical analysis of random mutagenesis methods used for directed protein evolution, J Mol Biol 355, 858-871. Zhang, N., Stewart, B. G., Moore, J. C., Greasham, R. L., Robinson, D. K., Buckland, B. C., and Lee, C. (2000) Directed evolution of toluene dioxygenase from Pseudomonas putida for improved selectivity toward cis-indandiol during indene bioconversion, Metab Eng 2, 339-348. Reetz, M. T., Torre, C., Eipper, A., Lohmer, R., Hermes, M., Brunner, B., Maichele, A., Bocola, M., Arand, M., Cronin, A., Genzel, Y., Archelas, A., and Furstoss, R. (2004) Enhancing the enantioselectivity of an epoxide hydrolase by directed evolution, Org Lett 6, 177-180. You, L., and Arnold, F. H. (1996) Directed evolution of subtilisin E in Bacillus subtilis to enhance total activity in aqueous dimethylformamide, Protein Eng 9, 7783.
192 [13] [14] [15]
[16]
[17] [18]
[19] [20] [21]
[22]
[23]
[24]
[25]
[26]
[27]
Birthe Borup, Lynne Gilson, Richard Fox et al. Xiang, Z. (2006) Advances in homology protein structure modeling, Curr Protein Pept Sci 7, 217-227. Wells, J. A., Vasser, M., and Powers, D. B. (1985) Cassette mutagenesis: an efficient method for generation of multiple mutations at defined sites, Gene 34, 315-323. Hayashi, N., Welschof, M., Zewe, M., Braunagel, M., Dubel, S., Breitling, F., and Little, M. (1994) Simultaneous mutagenesis of antibody CDR regions by overlap extension and PCR, Biotechniques 17, 310, 312, 314-315. Hibbert, E. G., Senussi, T., Costelloe, S. J., Lei, W., Smith, M. E., Ward, J. M., Hailes, H. C., and Dalby, P. A. (2007) Directed evolution of transketolase activity on non-phosphorylated substrates, Journal of biotechnology 131, 425-432. Reetz, M. T., and Carballeira, J. D. (2007) Iterative saturation mutagenesis (ISM) for rapid directed evolution of functional enzymes, Nature protocols 2, 891-903. Williams, G. J., Woodhall, T., Farnsworth, L. M., Nelson, A., and Berry, A. (2006) Creation of a pair of stereochemically complementary biocatalysts, J Am Chem Soc 128, 16238-16247. Giver, L., and Arnold, F. H. (1998) Combinatorial protein design by in vitro recombination, Curr Opin Chem Biol 2, 335-338. Stemmer, W. P. (1994) Rapid evolution of a protein in vitro by DNA shuffling, Nature 370, 389-391. Kurtzman, A., Govindarajan, S., Vahle, K., Jones, J., Heinrichs, V., and Patten, P. (2001) Advances in directed protein evolution by recursive genetic recombination: applications to therepeutic proteins, Curr Opin Biotech. 12, 361-370. Kawarasaki, Y., Griswold, K. E., Stevenson, J. D., Selzer, T., Benkovic, S. J., Iverson, B. L., and Georgiou, G. (2003) Enhanced crossover SCRATCHY: construction and high-throughput screening of a combinatorial library containing multiple non-homologous crossovers, Nucleic Acids Res 31, e126. Ryu, K., Hwang, S. Y., Kim, K. H., Kang, J. H., and Lee, E. K. (2008) Functionality improvement of fungal lignin peroxidase by DNA shuffling for 2,4-dichlorophenol degradability and H(2)O(2) stability, J Biotechnol 133, 110-115. Hsu, J. S., Yang, Y. B., Deng, C. H., Wei, C. L., Liaw, S. H., and Tsai, Y. C. (2004) Family shuffling of expandase genes to enhance substrate specificity for penicillin G, Appl Environ Microbiol 70, 6257-6263. Moore, J. C., Jin, H. M., Kuchner, O., and Arnold, F. H. (1997) Strategies for the in vitro evolution of protein function: enzyme evolution by random recombination of improved sequences, J Mol Biol 272, 336-347. Stemmer, W. P. (1994) DNA shuffling by random fragmentation and reassembly: in vitro recombination for molecular evolution, Proc Natl Acad Sci U S A 91, 1074710751. Stutzman-Engwall, K., Conlon, S., Fedechko, R., McArthur, H., Pekrun, K., Chen, Y., Jenne, S., La, C., Trinh, N., Kim, S., Zhang, Y. X., Fox, R., Gustafsson, C., and Krebber, A. (2005) Semi-synthetic DNA shuffling of aveC leads to improved industrial scale production of doramectin by Streptomyces avermitilis, Metab. Eng. 7, 27-37.
The Evolution of Directed Evolution [28] [29]
[30]
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Fox, R. (2005) Directed molecular evolution by machine learning and the influence of nonlinear interactions, J. Theor. Biol. 234, 187-199. Fox, R., Roy, A., Govindarajan, S., Minshull, J., Gustafsson, C., Jones, J., and Emig, R. (2003) Optimizing the search algorithm for protein engineering by directed evolution, Protein Eng. 16, 589-597. Fox, R. J., Davis, S. C., Mundorff, E. C., Newman, L. M., Gavrilovic, V., Ma, S. K., Chung, L. M., Ching, C., Tam, S., Muley, S., Grate, J., Gruber, J., Whitman, J. C., Sheldon, R. A., and Huisman, G. W. (2007) Improving catalytic function by ProSARdriven enzyme evolution, Nat Biotechnol 25, 338-344.
In: Biocatalysis Research Progress Editors: F. H. Romano, A. Russo
ISBN: 978-1-60456-619-2 © 2008 Nova Science Publishers, Inc.
Chapter VI
Biocatalytic Resolution of DL-Pantolactone by Cross-linked Cells and its Industrial Application Zhi-Hao Sun1, Ye Ni1, Pu Zheng1, Xin-Fu Guo2 and Jun Wang2 1
The Key Laboratory of Industrial Biotechnology, Ministry of Education; Laboratory of Biocatalysis, School of Biotechnology, Jiangnan University, 1800 Lihu Road, Wuxi 214122, PR China 2 Zhejiang Hangzhou Xinfu Pharmaceutical Co. Ltd., Hangzhou 311301, PR China
Abstract D-Pantoic acid (D-PA) and d-pantolactone (D-PL) are known as important intermediates for the production of calcium pantothenate, an additive for animal feed. Studies have showed that several filamentous fungi belonging to the genera Fusarium, Gibberella and Cylindrocarpon could produce D-PL hydrolases that selectively hydrolyze the D-isomer of DL-pantolactone to form D-PA,. In our previous studies, an excellent stereoselective D-lactonohydrolase producing strain Fusarium moniliforme CGMCC 0536 (SW-902) was obtained from isolation and mutation. Cells of this strain were successfully applied for the kinetic resolution of DL-pantolactone to produce D-(−)-isomer (99% e.e.). Furthermore, several immobilization methods were established to reuse the cells. The rationale for choosing whole cell immobilization is because it not only eliminates enzyme purification and extraction steps, but also increases enzyme thermostability, provides higher operational stability, greater resistance to environmental fluctuations and lower enzyme cost. An important technique is to treat cells with glutaraldehyde. The cross-linking of enzymes with glutaraldehyde involves the reaction between this bifunctional reagent and free amine groups of the enzyme. The linkages formed are irreversible and lead to cross-linked enzymes and
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cells exhibiting high operational stability. The wide utilization of glutaraldehyde for whole cells immobilization can be attributed to the decrease of enzyme leakage by cross-linking among the chemical reagent, cell wall, and intracellular protein. Therefore, we investigated the potential of cross-linking F. moniliforme CGMCC 0536 with glutaraldehyde for entrapping Dlactonohydrolase inside the cells and the employment of cross-linked biocatalyst in a stirred reactor. Our results have showed that cells of F. moniliforme CGMCC 0536 have been successfully immobilized by cross-linking with glutaraldehyde. The cross-linked cells exhibited a markedly improved thermal stability and operational stability than free cells. Kinetic characteristics of immobilized cells were assessed. The Km value of cross-linked cells (118 mM) was slightly higher than that of free cells (96 mM), while the Vmax value decreased from 4.18 to 3.87 mM min−1 g−1 wet cells after cross-linking. Furthermore, glutaraldehyde treatment did not change the stereospecificity, pH, and temperature profile of the D-pantonohydrolase. The high storage stability reduces fermentation workload. Significant process engineering advantages were evident for cross-linked cells in repeated batch operations. The resolution of DL-pantolactone was maintained at a steady level during 110 consecutive batches. The high activity and operational stability of the cross-linked cells presented in this work have been successfully implemented in the commercial production. Recently, we attempted to operate the resolution continuously by recycling cross-linked cells in a membrane bioreactor. The data show that glutaraldehyde cross-linking affords a satisfactory method for preserving the asymmetric hydrolyzing capacity of F. moniliforme CGMCC 0536. A feasible method with high operational stability for the production of D-PA catalyzed by crosslinked cells was established in a continuous process by using membrane bioreactor.
1. Introduction D-Pantoic acid (D-PA) and d-pantolactone (D-PL) are known as important intermediates for the production of calcium pantothenate, an additive for animal feed and various pharmaceutical products. Several derivatives of D-pantothenate, such as panthenyl alcohol, pantetheine, 4'-phosphopantetheine-S-sulfonate, coenzyme A, are also used for pharmaceutical products, as additives for infant formulae, and as chemical reagents. At present, the commercial production of D-pantothenate depends exclusively on chemical synthesis. Pantoic acid and pantolactone are conventionally produced by chemical methods, which yield an optically inactive racemic mixture that has to be resolved to obtain pure d-isomers. This process involves multiple reactions including yielding racemic pantolactone from isobutyraldehyde, formaldehyde and cyanide, optical resolution of the racemic pantolactone, and condensation of the D-pantolactone with β-alanine. Drawbacks of this chemical process are the troublesome resolution of the racemic pantolactone and racemization of the remaining L-isomer. In this resolution step, the use of an expensive alkaloid as a resolving agent is required. (Kataoka M et al. 1995)
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Concerning the food industry, the use of biocatalyst instead of expensive and toxic chemical agents for resolution of racemic pantolactone is both economical and friendly to the environment. This chapter gives a brief review on biocatalytic resolution of DL-pantolactone by crosslinked cells and its industrial application. A novel process for the production of calcium Dpantothenate and D-panthenol from d-pantolactone is also discussed in this article.
2. Biocatalytic Resolution of DL-pantolactone The various microbial enzymatic reactions such as stereospecific reduction of ketopantoyl lactone, (Hata H et al. 1987; Shimizu S et al. 1984, 1987) ketopantoic acid, (Kataoka M et al. 1990b) or 2’-ketopantothenate esters (Kataoka M et al. 1990a), stereoselective enzymatic oxidation and reduction system to produce d-PL from dlpantolactone (Shimizu S et al. 1987a; 1987b; Kataoka M et al. 1991), were reported. In these methods, there is a disadvantage of using expensive prochiral carbonyl compounds as starting materials. And the above processes have little practical significance because both the substrate concentration and reaction rate are low, and therefore D-PL of high optical purity are difficult to obtain. Additionally, redox reactions require co-factor/co-enzyme and stringent reaction conditions. There are also several reports on the application of enzymatic asymmetric hydrolysis for the optical resolution of pantoyl lactone (Glänzer et al. 1988; Bevinakatti and Newadkar 1989). In these cases, esterified substrates, such as O-acetyl- or O-formylpantoyl lactone, and lipases were used as starting materials and catalysts respectively. Since the lactonohydrolase hydrolyzes the intramolecular ester bond of pantoyl lactone, it is not necessary to modify the substrate (modified PL). This is one of the practical advantages of this enzyme. (Kataoka M. et al. 1995b) Racemic PL [or racemic pantoic acid (PA)] is still one of the cheapest and most available intermediates for D-PL synthesis; therefore, if the efficient reaction or enzyme for the stereospecific separation of unmodified racemic PL exists, it might be promising for practical purposes. (Kataoka M et al. 1996) A large number of investigations on the microbial resolution of DL-pantolactone to obtain D-PL have been reported. The process of complete degradation of L-PL in dlpantolactone is described in Japanese Examined Patent Application Publication No.19745/72 (Sakamoto K 1994a). However, half of the enantiomer substrate was wasted in this process. When racemic PL is used as a substrate for the hydrolysis reaction with stereospecific lactonohydrolase, only D- or L-PL might be converted to the corresponding acid (D- or LPA, respectively). If the racemic mixture could be resolved into L-PA with D-PL remained (Figure 1a), in the case of the L-PL-specific lactonohydrolase, the optical purity of the remaining D-PL might still be low, except that L-PL has been completely hydrolyzed. (Hua L et al. 2004) Alternatively, if the racemic mixture could be resolved into D-PA with the D-PL-specific lactonohydrolase (Figure 1b), D-PA of high optical purity could be constantly obtained regardless of the hydrolysis yield. The hydrolysate D-PA is easily separated from
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pantolactone with a solvent, and may then be converted to D-PL by heating under acidic conditions. The enzymatic resolution of racemic PL with D-PL-specific lactonohydrolase of F. oxysporum has been further studied (Kataoka M et al. 1995ab, 1996). It was reported that several filamentous fungi belonging to the genera Fusarium, Gibberella and Cylindrocarpon could produce D-PL hydrolase that selectively hydrolyze the D-isomer of DL-pantolactone to form D-PA. All the strains showed quite high hydrolysis activities and stereospecificities with high substrate concentrations (Table 1). Because these genera are closely related taxonomically, the enzyme responsible for the asymmetric hydrolysis in each strain might be similar.
Figure 1. The racemic PL is hydrolyzed with the stereospecific lactonohydrolase.
Table 1. Hydrolysis of DL-pantoyl lactone by the selected microorganisms (Kataoka M et al. 1995b) Hydrolysis rate (%) at 6h
10 h
16 h
Optical purity(% ee for D-pantoic acid)
Fusarium oxysporum AKU 3702 F. oxysporum AKU 3714 F. oxysporum AKU 3718 F. oxysporum IFO 5265 F. oxysporum IFO 30701 F. oxysporum IFO 31213 F. solani IFO 9955 F. semitectum IFO 30200 F. roseum IFO 30966 F. sp. 3-a Gibberella fujikuroi AKU3806
29.6 30.7 26.2 26.7 29.9 28.7 25.4 26.9 26.4 26.7
33.5 36.3 32.6 32.3 34.8 33.6 30.5 32.5 30.8 32.9
38.0 40.8 36.4 36.0 38.7 36.7 34.4 37.2 35.2 37.0
>93 >93 >93 >93 >93 >93 >93 >93 >93 >93
28.2
34.4
38.5
>93
Cylindrocarpon didymum AKU 3891
26.9
31.2
36.1
>93
Strain
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S. Shimizu’s group reported early that a fungus Fusarium oxysporum AKU 3702 produces a novel lactone-hydrolyzing enzyme that catalyzes reversible hydrolysis of various aldonate lactones. The enzyme was purified and characterized carefully, and shown to catalyze the stereoselective hydrolysis of D-pantoyl lactone (Shimizu et al. 1992). Furthermore, it was shown that the asymmetric hydrolysis ability of this enzyme could be used for the optical resolution of pantoyl lactone (Kataoka et al. 1995). This process has been applied for the practical production of D-pantoyl lactone, a chiral building block for the commercial production of D-pantothenic acid. Under the optimal conditions, D-PL in a racemic mixture (700 mg ml−1) was stereospecifically hydrolyzed to D-PA by F. oxysporum cells with automatic control of the pH of the reaction mixture at 7.0. The formation of L-PA was barely detected. After 24 h, the amount of PL hydrolyzed in the reaction mixture reached 322 mg ml−1 with an optical purity of 96% ee for D-PA (Kataoka M et al. 1995b). In previous studies, we obtained an excellent stereoselective D-lactonohydrolase producing strain, Fusarium moniliforme CGMCC 0536 (SW-902), through isolation and mutation. Cells of this strain were successfully applied for the kinetic resolution of DLpantolactone to produce D-isomer (99%ee) (Sun ZH 2001). The lactonase gene of Fusarium moniliforme for optical resolution of DL-pantoyl lactone was expressed in Saccharomyces cerevisiae. (Liu ZQ et al. 2004). And the D-pantonohydrolase of Fusarium moniliforme was selected for directed evolution through error-prone PCR combined with DNA shuffling for improved activity and pH stability using a convenient two-step high-throughput screening method based on the product formation and pH indicator. Compared with wild-type Dpantonohydrolase, the mutant exhibited a 10.5-fold higher specific activity; moreover, it could retain 85% of its original activity after incubation under low pH (at pH 6.0), while the wild type retained only 40% activity. (Liu ZQ et al. 2006) The high-level expression of a Fusarium lactonase gene in Aspergillus oryzae and the application of the recombinant microorganism for the optical resolution of racemic pantoyl lactone have also been performed. (Honda K et al. 2005).
3. Biocatalytic Resolution of DL-Pantolactone by Cross-linked Cells Although free cells of Fusarium moniliforme SW-902 possessed a higher lactonohydrolase activity, it was detected in preparative-scale reaction, that free cells could easily be broken by the shearing stress of the stirrer, leading to difficulties in recovery of cells by filtration and centrifugation. To resolve this problem, different methods have been tried, such as immobilization of whole cells. Some immobilization methods were established to reuse the cells (Tang YX et al. 2002). Most of successful immobilization methods employed polysaccharides such as κ-carrageenan, calcium alginate and gelatin as biocatalyst carriers. The whole cell immobilization was considered to be appropriate. The benefit for choosing whole cell immobilization is not only because it eliminates enzyme purification and extraction steps, but that it also increases enzyme thermostability, provides higher operational stability, greater resistance to environmental fluctuations and lower enzyme cost.
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The cells of F. moniliforme SW-902 were entrapped in κ-carrageenan. The immobilized cells exhibited a higher storage and operational stability than free cells, and were reused for as many as 30 times without any evident loss of lactonohydrolase activity. In the repeated batch stereospecific hydrolysis, a steady degree of hydrolysis pantolactone was obtained. After storing at 4 ºC for 8 weeks, the D-lactonohydrolase activity of the immobilized cells remained stable. The optimum temperature and optimum pH of enzymatic activity of the immobilized cells were the same as those of free cells of F. moniliforme SW-902 (55 ºC, pH 7.5). Studies on the stereospecific hydrolysis kinetics show that immobilized cells had a slightly higher Km value (151 mM) than free cells (113 mM). (Tang YX et al. 2002) S. Shimizu, et.al reported the practical hydrolysis of D-PL in a racemic mixture using F. oxysporum cells entrapped in calcium alginate gels as the catalyst. When the gels were incubated in 350 g l−1 DL-PL, 85%~93% of D-PL was hydrolyzed to D-PA of high optical purity (91%~95% ee). After 180 cycles (i.e. 180 days), the immobilized cells retained 60% of their initial activity. This enzymatic resolution has been in commercial operation since 1999. (Shimizu S et al. 2001; Sakamoto K et al. 2005) Zhang X et al (2007) also reported that immobilized lactonase of F. proliferatum ECU2002 performed quite well in repeated batch resolution of DL-pantolactone at a concentration of 35% (w/v), retaining 67% of initial activity after ten cycles of reaction and affording the product in 94~97% ee. Though the entrapment method using polymeric gel is simple and non-destructive, it results in a high concentration of carriers and a low concentration of biocatalysts within the matrix, leading to low specific activity and low volumetric productivity. Furthermore, fast diffusion of substrates and products through the entrapment system are required to obtain a high productivity. An alternative technique is to treat cells with glutaraldehyde. Table 2. Effect of various methods of immobilization on yield of activity (Tang YX et al. 2002) Methods Entrapment
Carriers Relative activity (%) κ-Carrageenan 59.1 Calcium alginate 50.0 Gelatin 35.9 Cross-linkage Glutaraldehyde 91.4 The relative activity is defined as enzyme activity ratio of the immobilized cells and free cells.
The cross-linking with glutaraldehyde was investigated. As shown in Table 2, the relative activity of lactonohydrolase of cross-linked cells was apparently higher than that of gel entrapped methods. Table 2 shows that cells of F. moniliforme CGMCC 0536 were successfully immobilized with glutaraldehyde. The relative residual activity of D-lactonohydrolase of cross-linked cells was apparently higher (91%) than that of gel entrapped methods (maximum 59%) because of the elimination of diffusion barrier across the gel. Glutaraldehyde is an excellent crosslinking agent that has been widely employed in the field of enzymes and cells immobilization. The cross-linking of enzymes with glutaraldehyde involves the reaction
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between this bifunctional reagent and free amine groups of the enzyme. The linkages formed are irreversible and lead to cross-linked enzymes and cells exhibiting high operational stability. Although it has been proposed that glutaraldehyde cross-links with enzymes via Schiff base, the mechanism of the reaction is not completely validated (Wong SS, Wong LJC, 1992). The wide application of glutaraldehyde with whole cells can be attributed to the decrease of enzyme leakage by cross-linking among the chemical reagent, cell wall, and parts of intracellular protein (i.e. immobilization in situ). Thus, it seemed of interest to access the potential of cross-linking F. moniliforme CGMCC 0536 cells with glutaraldehyde to entrap D-lactonohydrolase inside the cells and employing the cross-linked biocatalyst in a stirred reactor. Table 3. Effect of glutaraldehyde (GA) concentrations on enzyme activity GA concentrations (mM) 0
Relative activity (%) 100.0
5
95.7
10
92.9
15
88.6
20
84.3
25
75.7
30
68.6
The effect of the glutaraldehyde concentration on enzyme activity and operational stability of the biocatalyst was investigated (Table 3 and Figure 2). The treatment of cells with this reagent at concentrations between 5–30mM resulted in a significant increase in the half-life of the biocatalyst. However, high glutaraldehyde concentrations caused a noticeable decrease in D-pantonohydrolase activity at the same time. It was reported that the reaction with a low concentration of glutaraldehyde may facilitate intermolecular cross-linking and preserve native protein conformations, while high glutaraldehyde concentrations could promote intramolecular cross-linking, alter the structure of the active site and restrict substrate access (Payne JW, 1973). Using 20mM glutaraldehyde, both high enzyme activity and stable operation of immobilized cells were reached. Application of glutaraldehyde cross-linked cells led to high-yielding enzymatic resolution of DL-PL ( Hua L et al. 2004). However, batch processes suffer from disadvantages such as low efficiency because of frequent start up and shut down procedures, high labor costs, and batch-to-batch variations when the cross-linked cells were used for resolution of DL-racemic pantolactone at industrial scale. Another major disadvantage is the need to separate cross-linked cells at the end of each hydrolysis reaction through filtration, in which a marked loss of cells occurred, resulting in high processing costs. Consequently, this process is not practical for biocatalysts recycle in industrial application.
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Relative activity(%
100
50
0 0
2
4
6
8
10
Reuse times [GA]: 0mM (+); 5mM (□); l0mM (○); 15mM ( ); 20mM ( ); 30mM (×). The lactonohydrolase activity was determined in 0.5M Tris–HCl buffer (pH 7.5) for 1 h with gentle shaking. The relative activity is defined as the percentage of enzyme activity in the reuse to that in the first use. Figure 2. Operational stability of the cross-linked cells treated with different concentrations of glutaraldehyde (GA).
Table 4. Resolution of DL-PL by cross-linked cells in continuous process
No.
Total flow volume of substrate (L)
Final hydrolysis degree (%)
Average dilution rate (h−1)
Productivity of reactor (g/h·L)
1
1.5
32.3
0.14
4.61
2
3
30.1
0.18
5.47
3
7
29.9
0.16
4.65
Recently, we attempted to operate the resolution continuously using recycling crosslinked cells in a membrane bioreactor. For continuous operation, the biocatalysts must be separated from the reactants. Microfiltration and ultrafiltration have proven to be versatile separation processes for bioreactors based on the separation of biocatalysts by a semipermeable membrane, by which the economic feasibility of the process was also promoted. The entire production process was optimized by evaluating parameters such as concentrations, fluid velocity, pressure, reactor volume, and membrane surface. This continuous recycle membrane bioreactor is a time-saving and low-cost technology, as separation and reaction may be closely integrated, thus leading to continuous production and enzyme reuse. (Hua L et al. 2005) The data of the Figure 3 show that glutaraldehyde cross-linking affords a satisfactory method for preserving the asymmetric hydrolyzing capacity of F. moniliforme CGMCC 0536. A steady production of D-PA was maintained in a continuous process by using membrane
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25
600 500 400 300 200 100 0
Permeate flux (l h-1 m2 )
Accumulative D-PA in permeate (g)
bioreactor. In this closed reaction system, on-line separation of products from cells was achieved, unnecessary material waste was avoided, and the productivity was enhanced with reduced auxiliary operation time and energy cost. As shown in Table 4, the productivity in continuous operation was increased to 4.61~5.47 g / h·L, compared with 3.57 g/ h·L in repeated batch bioreactor operation with conventional filtration.
20 15 10 5 0 0
10
20
30
40
50
60
Time (h) Figure 3. Long-term resolution of DL-pantolactone and permeate flux for the membrane bioreactor. (∆) DPA in permeate; (○) permeate flux ( Hua L et al. 2005).
4. Industrial Application of Biocatalytic Resolution of DL-pantolactone by Cross-linked Cells As an important intermediate for various pharmaceutical products and additive for animal feeds, calcium D-pantothenic acid has broad applications and a huge market. The current world production of calcium D-pantothenate is about 12,000~14,000 tons per year. At present, the commercial production of D-pantothenate depends still on conventional chemical synthesis. Since 1999, the commercial enzymatic resolution of DL-PL has been initially performed in Japan. Shimizu S. et.al reported the practical enzymatic process using F. oxysporum cells entrapped in calcium alginate gels as the catalyst (Shimizu S et al, 1999a), and the immobilized cells were reused for 180 cycles. It has been shown that the new process is superior both economically and environmentally. (water−49%, CO2−30% and BOD−62%, compared with the traditional chemical resolution method) (Shimizu S et al. 2001, 2002; Kataoka M et al, 2007) In China, the enzymatic resolution of DL-PL on an industrial scale was also implemented using glutaraldehyde cross-linked cells. The use of immorbilization by cross-linking fungi mycelial cell walls without adding carrier, not only reduced immorbilization cost, but also avoided potential mass transport problems caused by the carrier. The resolution reaction time was decreased from 21~40 h to 3~5 h with an optical purity of 99% enantiomeric excess (ee)
Zhi-Hao Sun, Ye Ni, Pu Zheng, Xin-Fu Guo and Jun Wang
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for D-pantoic acid. The repeated batch resolution of DL-pantolactone using cross-linked cells and resting cells were shown in Figure 5. The cost of biocatalyst in large-scale production was greatly reduced by repeated use of cross-linked cells for over 180 times (Figure 5).
Hydrolysis degree(%)
Figure 4. Biocatalytic resolution of DL-pantolactone on a large scale of the 100,000L reactor by cross-linked whole-cell. 50 40 30 20 10 0
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Figure 5. The operational stability of cross-linked cells with glutaraldehyde (GA) on a large scale of biocatalytic resolution of DL-pantolactone.
Compared with conventional chemical resolution, the biocatalytic resolution for the production of calcium D-pantothenic acid and D-panthenol has remarkable advantages, including reduced raw material cost for 69.2%, reduced waste liquid and waste residue for 43.8%, reduced energy cost for 12.7%, and reduced manufacture cost for 26.5%. Importantly, the application of biocatalytic resolution method not only is environmental benign, but also results in improved product quality and product safety, and therefore, an advanced technology standard for the manufacture process has been established. The industrial application of biocatalytic resolution of DL-pantolactone by cross-linked cells of F. moniliforme CGMCC 0536 has been successfully commercialized for the production of calcium D-pantothenate on a large scale of 6,000 t/y at Xinfu Pharmaceutical Co. Ltd. (Hangzhou, Zhejiang, China). This cross-linked whole-cell bioprocess for preparing
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D-PL was scaled up for the production in the 100,000L reactor (the photograph of reactor equipment is shown in Figure 4).
References Bevinakatti HS, Newadkar RV. Lipase catalysis in organic solvents; transesterification of oformyl esters of secondary alcohols. Biotechnol. Lett, 1989, 11: 785-788. Bódalo A, Bastida J, Gómez JL, Gómez E, Alcaraz I, Asanza ML. Stabilization studies of laminoacylase- producing Pseudomonas sp. BA2 immobilized in calcium alginate gel. Enzyme Microb Technol, 1997, 21:64–69. Çetinus SA, Öztop HN. Immobilization of catalase into chemically crosslinked chitosan beads. Enzyme Microb Technol, 2003,32: 889–894. Chang MC, Tanaka J. XPS study for the microstructure development of hydroxyapatitecollagen nanocomposites cross-linked using glutaraldehyde. Biomaterials, 2002, 23: 3879–3885. Chibata I, Tosa T, Sato T. Immobilized aspartase-containing microbial cells: preparation and enzymatic properties. Appl Microbiol, 1974, 27: 878–885. Freeman A, Dror Y. Immobilization of “disguised” yeast in chemically cross-linked chitosan beads. Biotechnol Bioeng, 1994, 44: 1083–1088. Glänzer B I, Faber K, Griengl H. Microbial resolution of O-acetyl pantoyl lactone. Enzyme Microb. Technol. 1988, 10:689-690. Hata H, Shimizu S, Yamada H. Enzymatic production of D-(-)-pantoyl lactone from ketopantoyl lactone. Agric. Bid. Chem, 1987, 51: 3011-3016. Hendrickx M, Ooms C, Engels C, Van PE, Tobback P. Obstruction effect of carrageenan and gelatin on the diffusion of glucose. J Food Sci, 1987; 52: 1113–1114. Honda K, Tsuboi H, Minetoki T, Nose H, Sakamoto K, Kataoka M, Shimizu S. Expression of the Fusarium oxysporum lactonase gene in Aspergillus oryzae: molecular properties of the recombinant enzyme and its application. Appl Microbiol Biotechnol, 2005, 66: 520– 526. Hua L, Sun ZH, Leng Y, Hu ZQ. Continuous biocatalytic resolution of dl-pantolactone by cross-linked cells in a membrane bioreactor. Process Biochemistry 40 (2005) 1137–1142. Hua L, Sun ZH, Zheng P, Xu Y. Biocatalytic resolution of dl-pantolactone by glutaraldehyde cross-linked cells of Fusarium moniliforme CGMCC 0536. Enzyme Microb. Technol, 2004, 35: 161–166. Kataoka M, Hirakata M, Sakamoto K, Yamada H, Shimizu S, Optical resolution of racemic pantoic acid through microbial stereoselective lactonization in an organic solvent/water two-phase system. Enzyme Microb.Technol. 1996, 19: 307–310. Kataoka M, Honda K, Sakamoto K, Shimizu S. Microbial enzymes involved in lactone compound metabolism and their biotechnological applications. Appl Microbiol Biotechnol, 2007, 75: 257–266. Kataoka M, Shimizu K, Sakamoto K, Yamada H, Shimizu S. Lactonohydrolase-catalyzed optical resolution of pantoyl lactone: selection of a potent enzyme producer and
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optimization of culture and reaction condition for practical resolution. Appl Microbiol Biotechnol, 1995b;44: 333–338. Kataoka M, Shimizu K, Sakamoto K, Yamada H, Shimizu S. Optical resolution of racemic pantolactone with a novel fungal enzyme, lactonohydrolase. Appl Microbiol Biotechnol, 1995a; 43:974–977. Kataoka M, Shimizu S, Doi Y,Sakamoto K, YamadaH.Stereospecific reduction of ethyl 2’ketopantothenate to ethyl D-(+)-pantothenate with microbial cells as a catalyst. Appl. Environ. Microbiol, 1990, 56, 3595-3597. Kataoka M, Shimizu S, Yamada H. Stereospecific conversion of a racemic pantoyl lactone to D-(-)-pantoyl lactone through microbial oxidation and reduction reactions. Recl. Trav. Chim. Pays-Bas,1991,110, 155-157. Kataoka M, Shimizu S,Yamada H. Novel enzymatic production of D-(-)-pantoyl lactone through the stereospecific reduction of ketopantoic acid. Agric. Biol. Chem, 1990, 54, 177-182. Kennedy JF, Kalogerakis B, Cabral JMS. Immobilization of enzymes on cross-linked gelatin particles activated with various forms and complexes of titanium(IV) species. Enzyme Microb Technol ,1984, 6: 68–72. Liu ZQ, Sun ZH, Leng Y. Directed Evolution and Characterization of a Novel DPantonohydrolase from Fusarium moniliforme. J. Agric. Food Chem, 2006, 54: 58235830. Liu ZQ, Sun ZH. Cloning and Expression of D-Lactonohydrolase cDNA from Fusarium moniliforme in Saccharomyces cerevisiae, Biotechnology Letters, 2004, 26: 1861-1865. Payne JW. Polymerization of proteins with glutaraldehyde. Soluble molecular-weight markers. Biochem J, 1973;135: 867–873. Sakamoto K, Honda K, Wada K, Kita S, Tsuzaki K, Nose H, Kataoka M, Shimizu S. Practical resolution system for DL-pantoyl lactone using the lactonase from Fusarium oxysporum. J Biotechnol, 2005, 118: 99–106. Sakamoto K, Yamada H , Shimizu S. D-pantolactone hydrolase and process for the preparation thereof. US Patent, 5372940, 1994b. Sakamoto K, Yamada H , Shimizu S. Process for the Preparation of D-Pantolactone US Patent, 5275949, 1994a. Shimizu S, Hata H,Yamada H. Reduction of ketopantoyl lactone to &(-)-pantoyl lactone by microorganisms. Agric. Biol. Chem, 1984, 48: 2285-2291. Shimizu S, Hattori S, Ham H, Yamada H. One-step microbial conversion of a racemic mixture of pantoyl lactone to optically active D-(-)-pantoyl lactone. Appl. Environ. Microbiol, 1987, 53(3): 519-522. Shimizu S, Hattori S, Hata H, Yamada H. Stereoselective enzymatic oxidation and reduction system for the production of D(-)-pantoyl lactone from a racemic mixture of pantoyl lactone. Enzyme Microb. Technol, 1987, 9: 41l-416;. Shimizu S, Kataoka M, Chung MCM., Yamada H. Ketopantoic acid reductase of Pseudomonas mdtophilia 845, purification, characterization, and role in pantothenate biosynthesis. J. Biol. Chem, 1988, 263: 12077-12084.
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Shimizu S, Kataoka M, Honda K, Sakamoto K. Lactone-ring-cleaving enzymes of microorganisms: their diversity and applications. Journal of Biotechnology , 2001, 92: 187–194. Shimizu S, Kataoka M, Shimizu K, Hirakata M, Sakamoto K, Yamada H. Purification and characterization of a novel lactonohydrolase, catalyzing the hydrolysis of aldonate lactones and aromatic lactones, from Fusarium oxysporum. Eur. J. Biochem, 1992, 209: 383-390. Shimizu S, Morikawa T, Nitta K, Sakamoto K, Wada K. Biocatalytic Optical Resolution of DL-Pantolactone on an Industrial Scale. Journal of the Chemical Society of Japan, Chemistry and Industrial Chemistry (Japanese). 2002,(1):1-9. Shimizu S, Yamada H, Hata H, Morishita T, Akutsu S, and Kawamura M. Novel chemoenzymatic synthesis of n-(-)-pantoyl lactone. Agric. Biol. Chem, 1987, 51: 289290; . Sun ZH. A Method for Preparation of D-pantolactone by microorganisms Chinese Patent 01104070.X, 2001. Tang YX, Sun Z H, Hua L, Lv CF, Guo XF, Wang J. Kinetic resolution of dl-pantolactone by immobilized Fusarium moniliforme SW-902. Process Biochem, 2002,38: 545–549. Wong SS, Wong LJC. Chemical crosslinking and the stabilization of proteins and enzymes. Enzyme Microb Technol, 1992;14: 866–874. Zhang X, Xu JH, Xu Y , Pan J. Isolation and properties of a levo-lactonase from Fusarium proliferatum ECU2002: a robust biocatalyst for production of chiral lactones. Applied Microbiology and Biotechnology, 2007,75(5): 1087-1094.
In: Biocatalysis Research Progress Editors: F. H. Romano, A. Russo
ISBN: 978-1-60456-619-2 © 2008 Nova Science Publishers, Inc.
Chapter VII
Biocatalytic Potential of Haloalkaliphilic Bacteria Satya P. Singh*, Megha K. Purohit, Jignasha T. Thumar, Sandeep Pandey, Chirantan M. Raval and Hetal G. Bhimani Department of Biosciences; Saurashtra University, Rajkot- 360 005, India
Abstract Haloalkaliphilic organisms have been largely investigated from Soda lakes around the globe and other habitats for these organisms are rarely explored. During the last 10 years, we have been working on the diversity and enzymatic potential of haloalkaliphilic bacteria from the natural and man made saline habitats along the coastal Gujarat in Western India. The organisms have displayed varied diversity based on their cultural and morphological patterns, Gram reaction, biochemical properties, antibiotic resistancesensitivity, molecular phylogeny and secretion of extracellular enzymes. The production of extracellular alkaline proteases and lipases were widely spread among the isolates, whereas only few secreted amylase. The wide spread distribution of these bacteria from beyond soda lakes clearly indicated their ecological significance. Besides, the enzymatic potential would attract several biotechnological applications under alkalinity and high salt conditions. While major attention has been focused on molecular phylogeny and diversity, only limited information is available on their enzymatic potential and enzyme characterization. Cloning, sequencing and expression of different enzymes from Haloalkaliphilic bacteria and archaea are limited in literature. Our studies with alkaline proteases from a range of Haloalkaliphilic isolates revealed that many enzymes were highly resistant to urea denaturation and displayed catalytic potential under combination of extreme conditions. However, the ability to catalyse under extreme conditions was salt dependent. The
*
E- mail:
[email protected],
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Satya P. Singh, Megha K. Purohit, Jignasha T. Thumar et al. enzymes displayed unique features for biotechnological applications, besides providing a model system to study protein folding and stability. It is evident that the secretion and properties of alkaline proteases would also be very useful in assessment of microbial heterogeneity. The prospects of metagenomics to explore the novel sequences for biocatalysts from non-cultivable microbes have emerged as a potential tool in recent years. This would also be discussed with particular reference to saline habitats. Over all, our findings would be integrated with the literature and biocatalytic potential of Haloalkaliphilic bacteria and archaea would be assessed.
Keywords: Haloalkaliphilic bacteria, thermostability, alkaline proteases, non-aqueous biocatalysis, metagenomics, Cloning & expression of salt-tolerant enzymes
Introduction Extremophiles are unique microorganisms that can grow and thrive in extreme conditions such as; temperature, pressure, radiation, desiccation, salinity, pH, oxygen species, redox potential, metals and gases (van den Burg, 2003; Javaux, 2006). Among the extremophiles, halophiles constitute a rather heterogeneous group (Ventosa et al., 1998; Oren, 2002(a) and 2002(b) and are primarily categorized on the basis of their optimum salt requirements; extreme halophiles (15–30%, w/v) and moderate halophiles (3–15%, w/v) (Kushner, 1978; Ventosa et al., 1998). Extreme halophiles accumulate intracellular K+ (Kushner, 1978; Oren, 1999), while moderate halophiles preferentially accumulate compatible solutes to cope with the osmotic stress in their habitats (Oren, 1999). Haloalkaliphilic bacteria, able to grow under high salt and alkaline pH, have largely been explored and studied from the concentrated hyper saline environments - Soda Lake, Solar Saltern, Salt brines, Carbonate springs and Dead Sea. Where as, the exploration of the natural saline and alkaline environments beyond the above boundaries is just the beginning. Our work on these bacteria over the last several years has indicated their wide occurrence in the habitats beyond Soda lakes. The Haloalkaliphilic bacteria were isolated under different enrichment conditions and characterized on the basis of their colony and cell morphology, Gram reaction, biochemical properties, antibiotic resistance-sensitivity, plasmid profiling and 16S rRNA gene sequence homology (Thumar and Singh 2007(b); Dodia et al., 2006; Nowlan el al., 2006; Parmar et al., 2006,Vasavada et al., 2006) During the last few years, microbes with more than one degree of extremity have focused greater attention due to their biotechnological and ecological significance. Such microbes are being explored from their diversity, molecular phylogeny and biotechnological standpoint.
Biocatalytic Potential of Haloalakliphilic Bacteria The increasing emphasis on multi-extremophiles is due to the fact that the "survival kits" of these microbes can potentially serve in an array of applications. In addition, these organisms may hold secrete for the origin of life and stability of the macromolecules. Of
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particular interest are the enzymes (biological catalysts) that help multi-extremophiles to function in brutal circumstances. They can, therefore, form the basis of entirely new enzymebased processes. The industrial applications of enzymes able to withstand harsh conditions have greatly increased over the past decade. This is due to the discovery of novel enzymes from extremophilic microorganisms. Recent advances in the study of extremozymes point to the acceleration of this trend (Demirjian et al., 2007). In particular, enzymes from thermophilic and halophilic organisms have found the most practical commercial uses to date because of their overall inherent stability. This has also led to a greater understanding of stability factors involved in adaptation of these enzymes to their unusual environments (Burg, 2003). Most of the studies on Haloalkaliphilic bacteria, however, have so far focused on phylogenetic analysis of the organisms and only limited information is available on their enzymatic and other biotechnological potential. Our studies have clearly indicated wide occurrence of the extracellular enzymes, alkaline proteases, in particular (Thumar and Singh 2007 (a), Dodia et al., 2007(a)and 2007(b); Patel et al., 2006(a) and 2006(b)., Gupta et al., 2005., Parmar et al., Sinha et al., 2007 ) The haloalkaliphilic strains from sea water and other saline habitats along coastal Gujarat secreted most commonly occurring hydrolytic and polymer degrading enzymes; proteases, amylases, chitinases and lipases. The seawater isolates had wide occurrence and variation in production of alkaline proteases, chitinases and lipase, where as only few secreted amylase. The pattern suggested that occurrence of extracellular enzymes could also be used as biochemical marker to judge the microbial heterogeneity among moderately haloalkaliphilic bacteria. Besides, dual extremities of alkaline pH and salinity coupled with high temperature stability of enzymes in many cases projected them as promising candidates for various biotechnological applications. The proteases were thermally stable where salt acted as positive effector. In addition, majority of the enzymes displayed salt-dependent resistance against chemical denaturation, a feature which is quite rare among the proteins. The alkaline proteases from a number of haloalkaliphilic bacteria isolated from Coastal saline habitats of Gujarat in Western India have displayed unique features for biotechnological applications, besides providing a model system to study protein folding and stability under extreme set of conditions (Dodia et al., 2007a and 2007b). The advances in molecular tools and computational biology would add impetus in search, screening and evolution of new generation of biocatalysts for novel applications.
Cloning and Expression of Enzymes from Haloalkaliphilic Bacteria Among the enzymes from extremophilic organisms, relatively limited awareness exists about enzymes from haloalkaliphilic bacteria. Maintenance of stability and activity in high salt is major challenge for halophilic proteins. Most typical halophilic enzymes from extremely halophilic archaea and bacteria require high concentrations of salt for their activity and stability and are inactivated in Escherichia coli unless refolded in the presence of salts
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under in-vitro conditions. Recombinant DNA Technology in conjunction with many other molecular techniques is being used to improve and evolve enzymes leading to new opportunities for biocatalysis. Therefore, cloning of the potential genes coding for different enzymes would be an attractive approve to begin with. The gene encoding a ferredoxin of nucleoside diphosphate kinase from a moderately halophilic eubacterium was cloned and protein was over expressed in E.coli. Sequence analysis of the cloned gene revealed an open reading frame of 387 nucleotides encoding 129 amino acids. The deduced amino acid sequence of H. Japonica Fd showed 84 to 98% identity with the corresponding sequences of other extremely halophilic archaea (Matsuo et al., 2001). The extracellular-amylase-encoding amyH gene isolated from a moderate halophile, is understood to be the first extracellular-amylase with significant biotechnological potential. Besides, H. meridiana and H. elongata were also able to secrete the thermostable -amylase from Bacillus licheniformis, indicating that members of the genus Halomonas could be good candidates for the production of heterologous extracellular enzymes (Bautista et al., 2006). Some alkaline protease-encoding bacterial genes have been cloned and expressed in new hosts, the two major organisms for cloning and over-expression being E. coli and B. subtilis. The gene of a highly thermostable alkaline protease from an alkaliphilic bacillus was cloned by PCR and nucleotide sequence was determined. Similarly, around 1242 base pair DNA fragment from B. halodurans isolated from alkaline sediments coding for a potential protease was cloned and sequenced. As deduced from amino acid sequence, it was an active monomer of 46.5 kDa (Calik et al., 2003). This recombinant F1 protease was efficiently secreted into the culture medium using E. coli harboring two vectors with its lac promoter–operator system. Another alkaline proteinase (subtilisin) gene was cloned and sequenced from alkaliphilic B. lentus NCIB 10309 into B. subtilis DN497 (Jorgensen et al., 2000). A new strain of B. pumilis, c172-14 (pBX96) was engineered by introducing the pBX 96 plasmid (carrying the α-amylase amy gene) into the host strain of alkalophilic B. pumilis c172 through transformation. The level of alkaline protease production was improved to 43% of new strain compared to the parent strain (Feng et al., 2001). Another alkaline protease gene, apr, from B. licheniformis 2709 was cloned into a Bacillus shuttle expression vector, pHL and expressed in Bacillus subtilis WB600. The expression of alkaline protease increased by 65% in the engineered strain BW-016 relative to the original strain (Tang et al., 2004). Recently, apr 46 gene encoding protease was cloned from Bacillus licheniformis RSP-09-37 and functionally expressed in E. coli JM 109. The recombinant protease displayed 29-50% homology with known serine proteases and conserved domains (Sareen et al., 2005). Earlier, gene encoding alkaline serine protease (Halolysin) was cloned and expressed in Haloferax volcanii from an unidentified halophilic archaeal strain (172 PI) by Kamekura et al. (1992). The cloning, sequencing, and specific amplification of a protease gene on the chromosome of an alkaliphilic bacillus was carried out with increase in gene copy number by an improved gene amplification technique. Further, the gene encoding ferric uptake regulator protein (fur gene) of Vibrio (2088bp fragment), encoding a protein of 147 amino acids, and homologous with fur, was identified, cloned and sequenced. The fur gene was subsequently amplified and comparison of phylogenetic analyses using fur and 16S DNA coding for rRNA sequences, confirmed the
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usefulness of fur as an evolutionary marker (Colcohoun, 2003). Earlier, to study gene expression in halophilic archaea, a reporter system was analyzed by β–glycosidase enzyme. The enzyme was active at 4 M NaCl and consisted of two monomers with estimated molecular weight of 180±20 kDa (Jolley et al., 1996). The developments related to cloning and expression of the genes from halophilic organisms in heterologous hosts will certainly boost the number of enzyme-driven transformations in chemical, food, pharmaceutical and other industrial applications.
Metagenomic Approaches towards Exploring Biocatalytic Potential of Halophilic and Haloalkaliphilic Organisms Metagenomics, an emerging approach to explore diversity and harness microbial potential, is based on the analysis of the genomic DNA of microbial communities in their natural environments and cloning it into a cultured organism (Danchin et al., 2004). Advances have derived from both sequence-based and functional analysis of the metagenome. Studies on metagenomes have revealed the vast scope of biodiversity in a wide range of environments, new functional capacities of individual cells and communities and their complex evolutionary relationships (Rondon, 2000). The continuous advancement in the metagenomics has dramatically revised our view of microbial biodiversity and its potential for biotechnological applications. In the light of the estimation that 95-99% of microorganisms in most environments are not cultivable, quite less is known about their genomes, genes and encoded enzymatic activities. The isolation and analysis of environmental DNA has enabled us to mine microbial diversity, allowing us to access their genomes, identify protein coding sequences and even to reconstruct biochemical pathways towards understanding the properties and functions of these organisms. The metagenomic libraries would, thus, provide a huge genetic resource base. However, with respect to extreme habitats, the extremophilic metagenome is the least explored and understood. Though the complete metagenome of the microbial community of a crystallizer pond has been analyzed by end sequencing a 2000 clone fosmid library and compared with the genome sequence of Haloquadratum walsbyi (Legault et al., 2006). The environmental DNA library was completely retrieved, but many ORF’s ascribed to the Haloquadratum meta-population by common genome characteristics or scaffolding to the strain genome were not present in the specific sequenced isolate. The extensive gene repertoire is expected of a population as diverse nutrient pool is available, resulting in highly divergent population. 16S rRNA-based phylogenetic profiling of hyperthermophiles samples from various geothermal sites are also studied through metagenomics to get insights into the community structure and to relate metabolic characteristics in high-temperature habitats. Genes linked to environmental functionality with novel archaeal lineages having unusual 16S rRNA and protein-coding ability were identified from biofilms at pH 0.5-1.5 in acid mines through metageniomic studies (Raes et al., 2007). The Comprehensive knowledge of the genetic blueprints, the functions and the interactions of microbial communities will provide
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insight into the evolution of marine ecosystems. Our research group at the Saurashtra University is engaged with halophilic/haloalkaliphilic bacteria from coastal region of Gujarat (Western India) for the last several years and one of the current focuses is the metagenomic approach to study diversity and biocatalytic potential of the non-cultivable organisms from these saline habitats. The advances in metagenomics would revolutionize the investigations on microbial ecology and biotechnology, leading to exploration of uncultured microbial population and discovery of new enzymes for various applications (Scheimmezer et al., 2007). The successful cloning of a novel decarboxylase gene was achieved using metagenomic library genes from alkaline polluted soils. The recombinant protein was over expressed and functional characterization was followed by liquid chromatography-mass spectrometry. The particular areas of interest relate to randomly proliferating limited-source DNA, parallel sequencing without cloning, isolating specific target sequences from highly complex template mixtures, high-throughput assays for metabolic pathways, artificial transcriptional regulators activating reporter genes to indicate enzymatic substrate conversion and cDNA cloning from extracted mRNA from a microbial consortium. While the potential are huge, the task is not unchallenged. Among the uphill tasks, unavailability of the efficient heterologous expression systems for obtaining potential enzymes from unknown source organisms deserves worth mention.
Halophilic, Haloalkaliphilic Bacteria and their Enzymes in Organic Solvent Organic solvents are toxic to microorganism and the index of biological toxicity is estimated by log Pow (common logarithm of partition coefficient of given solvent in a mixture of n-octanol and water). A lower value of log Pow signifies higher toxicity (Inoue et al., 1991). Focus on the tolerance of microorganism and their enzymes to organic solvent gained momentum since the discovery of toluene-tolerant Pseudomonas putida strain IH-2000 (Inoue et al., 1989). A halophilic archaeal strain EH4 (Bertrand et al., 1990, Ward et al., 1978, Oren et al., 1992, Carla et al., 2004) was capable of degrading a wide range of nalkanes and aromatic hydrocarbon in the presence of high salt. In 1991 (Kulichevskaya et al., 1991) and 1995 (Zvyagintseva et al., 1995), few halophilic strains were isolated from a plot of Kalamkass oil field filled with saturated brine and oil. In addition, Marrinobacter hydrocarbonclasticus degraded a variety of aliphatic and aromatic hydrocarbons (Gauthier et al., 1992). A halotolerant streptomyces sp. isolated from oil field in Russia, was capable of degrading crude petroleum (Kuznetnov et al., 1992). In 2003, solvent tolerance by 16 genera of Halobacteriace (halophilic Archaea) was published (Usami, 2003). According to this study, strains were capable to grow in the presence of solvents with log Pow above 4.9. In the presence of organic solvents, two growth patterns were observed; in pattern A, the growth rate at exponential phase was considerably lower in presence of organic solvent but final density reached above 83% of control while in pattern B; both, growth rate and the final cell density were low. Halophilic bacteria isolated from Seminol County in Oklahoma, USA, referred as Sem-2, used benzene as sole carbon source (Carala et al., 2004). It also degraded
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toluene, ethylbenzene and xylenes as sole carbon source. Among all tested compounds, toluene was most favored. Solvent tolerance of a halophilic extracellular α-amylase has been recently reported (Tadamasa et al., 2005). The enzyme was produced by Haloarchaeon, Haloarcula sp. strain S-1. This amylase exhibited maximal activity at 50°C in 4.3 M NaCl, pH 7. Similarly, an extracellular protease produced by haloalkaliphilic archaeon Natrialba magadii (Diego et al., 2006) was active and stable in aqueous-organic solvent mixture containing 1.5 M NaCl and glycerol, Dimethylsulhooxide (DMSO), N,N-dimethyl formamide, propylenglycerol. Protease was more efficient at pH 8 than at pH 10. A metlloprotease from moderate halophilic bacterium Salinivibrio sp. strain AF-2004 also exhibited tolerance against organic solvents (Reza et al., 2006). The optimum temperature and salinity of the enzyme were at 55°C and 0-0.5 M NaCl, although at salinities up to 4M NaCl, the enzyme was still active. The protease was stable and had a broad pH profile (5.0-10.0) with an optimum 8.5 for casein hydrolysis. The application of alkaline proteases and other enzymes is well documented and its operation in organic media is an interesting area for research and applications in biotechnology. Generally, alkaline proteases are thermally stable in the range of 37-70°C. Alkaline proteases from haloalkaliphilic organisms in biphasic medium have been studied in only limited sense. Large number of Haloalkaliphilic bacterial isolates obtained from saline habitats of coastal Gujarat in western India, were able to grow and secret enzyme in the presence of different organic solvents such as butanol, methanol, n-hexane and propanal. The findings are indicative of future applications of these biocatalysts. The stability of the alkaline proteases in the presence of organic solvent would be an attractive feature of the biocatalysis.
Future Perspective The larger umbrella of genomics and proteomics has yet to cover haloalkaliphiles, where genomic information will provide deeper molecular insights into the ways microorganisms cope with more than one extremity. The increasing attention on these microbes is reflected by the regularly held conferences and symposia in various parts of the world indicating wealth of informations. Among the dimensions of further explorations; molecular phylogeny, structural basis of protein stability under extreme conditions, development of expression systems, over-expression and protein folding under In-vitro conditions are key to mention.
Acknowledgemnet The work highlighted from our own group was sponsored by Saurashtra University and UGC, Government of India. Contributions of Drs. Rajesh Patel, Mital Dodia and Rupal Joshi to our research programme, as doctoral students are also acknowledged. The further work on bio-cataloguing, molecular phylogeny and development of recombinant enzymes from halophilic and haloalkaliphilic bacteria of Coastal Gujarat in Western India are being
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sponsored by a DBT (Government of India): multi-institutional project (Saurashtra University-Rajkot, IIT- Delhi and Delhi University South Campus).
Refrences Bautista, V., Esclapez, J., Espinosa, R., Pomares, F.,Camacho, M. & Bonete, M. (2006). Heterologous overexpression of a halophilic α-amylase. Microbial Cell Factories., (Suppl 1):P13doi:10.1186/1475-2859-5-S1-P13. Betrand, J., Acquaviva, M. & Mille, G. (1990). Biodegradation of hydrocarbons by an extremely halophilic archaebacterium. Lett Appl. Microbiol. 11, 260-263. Burg, B. (2003). Extremophiles as a source for novel enzymes. Current Opinion in Microbiology, 6(3), 213-218. Çalık, P., Kalender, N. & Tunçer, H. (2003). Overexpression of serine alkaline protease encoding gene in Bacillus species: performance analyses. Enzyme & Microbial Technology, 32, 706–720. Carala, A., Nicholson & Babu, Z. (2004). Biodegradation of Benezene by halophilic & halotolerant bacteria under aerobic conditions. Applied & environmental Microbiology, 1222-1225. Colquhouna, D. & Sorumb, H. (2002). Cloning, characterisation & phylogenetic analysis of the fur gene in Vibrio salmonicida and Vibrio logei. Gene, 296, 213–220. Danchin, A. (2004). Microbial Cell Factories.3:13 doi: 10.1186/1475-2859-3-13. Demirjian, D., Varas, F. & Cassidy, C. (2001). Enzymes from Extremophiles. Current Opinion in Chemical Biology , 5(2),144-151. Diego, M. R., Rosana, E. De castro. (2006). Effect of organic solvents on the activity & stability of an extracellular protease secreted by haloalkiphilic archaeon Natrialba magadii. J Ind Microbiol Biotechnol. 23(4): 1432-1437. Dodia, M., Bhimani, H., Rawal C., Joshi, R. & Singh, SP. (2007b). Salt dependent resistance against chemical denaturation of alkaline protease from a newly isolated Haloalkaliphilic Bacillus sp. Bioresource Technology (In Press). Dodia, M., Joshi, R., Patel, R. & Singh, SP. (2006). Characterization and Stability of Extracellular Alkaline Proteases from Moderately Halophilic and alkaliphilic Bacteria Isolated from Saline Habitat of Coastal Gujarat, India. Brazilian Journal of Microbiology, 37, 276-282. Dodia, M., Rawal, C., Bhimani, H., Joshi R. H., Khare, S. & Singh,S. (2007a). Purification and stability characteristics of an alkaline serine protease from a newly isolated Haloalkaliphilic bacterium sp. AH-6 Journal of Industrial Microbiology & Biotecnology (In Press). Feng, Y., Yang, W., Ong, S., Hu, J. & Ng, W. (2001). Fermentation of starch for enhanced alkaline protease production by constructing an alkalophilic Bacillus pumilus strain. Appl Microbiol. Biotechnol. , 57(2),153-160. Fukushima, T, Mizuki, T., Echigo, A., & Usami, R. (2005). Organic solvent tolerance of Halophilic α-amylase from Haloarchaeon, Haloarcula sp. strain S-1. Extremohiles, 9, 8589.
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Gauthier, M. J., B. Lafay, R. Christen, L. Fernandez, M. Acquaviva, Bonoi. & J.C. Bertand. (1992). Marrinobacter hydrocarbonclasticus gen. nov., sp. Nov., a new extremely halotolerant, hydrocarbon-degrading marine bacterium. Int. J. Syst. Bacteriol. 42, 568576. Gupta, A., Roy, I., Patel, R., Singh, SP., Khare, S. & Gupta, M.N.(2005).One-step purification and characterization of an alkaline protease from Haloalkaliphilic Bacillus sp. Journal of Chromatography A, 1075, 103-108. Hamid, R., Karbalaei H., Ziaee, A. & Amoozegar, M. A. (2006). Purification & biochemical characterization of protease secreted by the Salinivibrio sp. strain AF-2004 & its behavior in organic solvents. Extremophiles, 11,237-243. Inoue, A. & Horikoshi, K. (1991). Estimation of solvent-tolerance of bacteria by solvent parameter log P. J.Ferment. bioeng., 71, 194-196. Inoue, A., & Horikoshi, K. (1989). A Psedomonas thrives in high concentration of toluene. Nature, 338, 264-266. Jolley, K., Rapaport, E., Hough, D., Danson, M., Woods, W. & Smith, D. (1996). Dihydrolipoamide Dehydrogenase from the Halophilic Archaeon Haloferax volcanii: Homologous Overexpression of the Cloned Gene. Journal of Bacteriology. 178(11), 3044–3048. Kamekura, M., Seno, Y., Holmes, M. L. & Dyall-Smith, M.L. (1992). Molecular cloning & sequencing of the gene for a halophilic alkaline serine protease (halolysin) from an unidentified halophilic archaea strain (172P1) & expression of the gene in Haloferax volcanii. J Bacteriol., 174(3),736-42. Kulichevskaya, I. S., Milekhina, E. I., Borzenkov, I. A., Zvyagintseva, I.S. & Belyaev. (1991). Oxidation of petroliumhydrocarbons by extremely extremely halophilic archaebacteria. Mikrobiologya, 60, 860-866. Kuznetnov, V. D., Zaitseva, T. A, Vakulenko. L. V. & Filippiva, S. N. (1992). Streptomyces albiaxialis Sp. nov.- a new petroleum hydrocarbon degrading species of thermo & halotolerant Streptomyces. Mykrobiologiya, 61, 84-91. Legault, B., Lopez, A., Carlos, J., Doolittle, W., Bolhuis, H., Rodriguehz, F. & Papke, T. (2006). Environmental genomics of "Haloquadratum walsbyi" in a saltern crystallizer indicates a large pool of accessory genes in an otherwise coherent species. BMC Genomics, 7 (1):171-174. Matsuo, T., Ikeda, A., Seki, H., Ichimata, T., Sugimori, D. & Nakamura, S. (2001). Cloning & expression of the ferredoxin gene from extremely halophilic archaeon Haloarcula japonica strain TR-1. BioMetals, 14,135–142. Nowlan B., Dodia, M.S., Singh, SP & Patel, B. K. C. (2006). Bacillus okhensis sp. nov., a halotolerant and alkalitolerant bacterium from an Indian saltpan. Int J Syst Evol. Microbiol., 56, 1073-1077. Nowlan, B., Dodia, M., Singh, SP., & Patel, B. (2006). Bacillus okhensis sp. nov., a halotolerant and alkalitolerant bacterium from an Indian saltpan. Int J Syst Evol. Microbiol., 56, 1073-1077. Oren, P., Gurevich, M., Azachi, & Y. Henis . (1992). Microbial degradation of pollutants at high salt concentration. Biodegradation, 3, 387-398.
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Parmar, K., Dodia, M., Joshi, R. & Singh., SP. (2006). Degradation of Azo Dyes by Amylase Secreting Haloalkaliphilic Bacteria Isolated from Saline Habitats in Mithapur and Veraval of Gujarat. Pollution Research, 25 (4), 1-5. Patel, R., Dodia, M. & Singh, SP. (2006b). Purification and characterization of alkaline protease from a newly isolated Haloalkaliphilic Bacillus sp. Process Biochemistry, 41(9), 2002-2009. Patel, R., Dodia, M., Joshi, R. & Singh, SP. (2006a). Production of extracellular haloalkaline protease from a newly isolated Haloalkaliphilic Bacillus sp. isolated from seawater in Western India. World Journal of Microbiology & Biotechnology, 22(4), 375382. Raes, M. (2007). Prediction of effective genome size in metagenomic samples. Genome Biology, 8:R10. Ron, Usami., Tadamasa, Fukushima., Toru, Mizuki., Akira, Inoue., Yasuhiko, yoshida. & Koki, Horikoshi. (2003) Organic solvent Tolerance of Halophilic archaea. Biosci. Biotechnol. Biochem., 67 (8), 1809-1812. Rondon M., August P., Bettermann D., Brady S., Grossman T., Liles A., Loiacano K., Lynch B., Minor C., Tiaong, C., Clardy, J. & Handelsmann, J. (2000). Cloning the Soil Metagenome a Strategy for Accessing the Genetic & Functional Diversity of Uncultured Microorganisms. BMC Genomics, 66(6), 2541–2547. Sareen, R., Bornscheuer, U.T. & Mishra, P. (2005). Cloning, functional expression & characterization of an alkaline protease from Bacillus licheniformis. Biotechnol Lett., 27 (23-24), 1901-1907. Schmeisser, C. & Stelle, S. (2001). Metagenomics, biotechnology with non-culturable microbes. Applied Microbiology & Biotechnology. 75(5), 356-364. Sinha, R., Joshi, R., Dodia, M. & Singh, SP. (2007). Production, purification and characterization of an alkaline protease from an alkaliphilic bacillus sp. Journal of Cell and Tissue Research, 7 (2), 1031-1037. Tang, X.M., Lakay, F.M., Shen, W., Shao, W.L., Fang, H.Y., Prior, B.A., Wang, Z.X. & Zhuge, J. (2004). Purification and characterization of an alkaline protease used in tannery industry from Bacillus licheniformis. Biotechnol Lett., 26 (18),1421-1424. Thumar, J. & Singh, S. (2007b). Secretion of an alkaline protease from a salt- tolerant and alkaliphilic, Streptomyces clavuligerus strain Mit-1. Brazilian Journal of Microbiology, (Accepted). Thumar, J. & Singh, S.P. (2007a). Two - step purification of a highly thermostable alkaline protease from salt-tolerant alkaliphilic Streptomyces clavuligerus strain Mit-1. Journal of Chromatography B. 854, 198–203. Vasavada, S. Thumar, J. & Singh, SP. (2006). Secretion of a potent antibiotic by salt-tolerant and alkaliphilic actinomycete Streptomyces sannanensis strain RJT-1. Current Science, 91 (10), 1393-1397. Ward, D.M. & T. D. Brock.(1978). Hydrocarbon biodegradation in hyper saline environments. Appl. Environ. Microbiol. 35, 353-359. Zvyagintseva, I.S., Belyaev, S., Borzenkov, I. A., Kostrikina, N.A., Milekhina, E. I. & Ivnov, M. V. (1995). Halophilic archaebacterium from Kalamkass oil field. Mikrobiologya, 64, 83-87.
In: Biocatalysis Research Progress Editors: F. H. Romano, A. Russo
ISBN: 978-1-60456-619-2 © 2008 Nova Science Publishers, Inc.
Chapter VIII
Salt-tolerant Alkaliphilic Actinomycetes and their Biocatalytic Potential Satya P. Singh* and Jignasha T. Thumar Department of Biosciences; Saurashtra University, Rajkot- 360 005, India
Abstract Extremophiles are distributed over a range of extreme habitats. Among them, extremophilic actinomycetes have recently attracted greater attention due to their various natural products and specific mechanism of adapting extreme environments. The natural and man-made environments may harbor a large population of halophilic and alkaliphilic actinomycetes. However, they have only recently focused attention of the researchers. The phylogeny, diversity and biotechnological potential of salt-tolerant alkaliphilic actinomycetes are still in infancy. It is, therefore, relevant and important to pay more attention to extreme actinomycetes from unexplored habitats, as a possible way to discover novel taxa and, consequently, new secondary metabolites. The study would also enlighten us on their diversity, phylogeny and ecological significance. During the last decade, there has been a dramatic increase in the need for bioactive compounds with novel activities. Enzymes, after antibiotics are the most important biologically derived product having immense potential in catalytic reactions of commercial interest. Most of the studies related to enzymes have so far focused on halophiles, alkaliphiles and haloalkaliphiles; however, the enzymatic potential of halotolerant alkaliphilic actinomycete is nearly untouched. From the literature, it is evident that the exploration of the enzymatic potential of these microbes is just the beginning and till date only few enzymes are investigated in depth. Salt-tolerant alkaliphilic actinomycetes produce enzymes, such as alkaline protease, amylase, cellulase and lipase that are functional under extreme conditions. Consequently, the unique properties of these biocatalysts have potentials in several novel applications in industrial processes.
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Satya P. Singh and Jignasha T. Thumar Because of the capacity to survive under nonstandard conditions in non-conventional environments, it is assumed that the properties of extremophilic enzymes have been optimized for these conditions. Interestingly, our own studies have also revealed the occurrence of alkaline protease, amylase and cellulase in salt-tolerant alkaliphilic actinomycetes isolated from vast coastal line and saline habitats along the Saurashtra Coast Gujarat (Western India). In addition, the results on the alkaline proteases are suggestive of their unique position in the generation of novel enzymes.
Keywords: Salt-tolerant alkaliphilic actinomycetes, extremophilic enzymes, biocatalytic potential, alkaline proteases
Introduction Microorganisms have traditionally played a major role in biotechnology exhibiting range of metabolic diversity and habitat preferences not observed in higher organisms. However, only fraction of microbial world has been explored due to our inability to grow majority of them into the laboratory. This fact assumes further significance with regards to microbes from extreme environments as such ecological niches are only poorly investigated for the resident microbial flora and their ecological significance. Therefore, exploring more and more such extreme habitats would be of particular interest. During the long history of human beings, the most concentrated and widespread occurrences of organisms are generally observed in “moderate” environments. It has also been known that certain environments on earth are thought to prevent the existence of life (Horikoshi et al., 1991). At different times, microorganisms have been isolated from the different environments, including extreme ones. The classification of “extreme environments” refers to a wide variety of different conditions to which microorganisms have adapted. It can not only refer to chemical extremes e.g. salinity, pH and toxic chemicals, but also to physical extremes such as temperature and pressure. Organisms which thrive in extreme environments offer the opportunity to appreciate the range of adaptive possibilities on fundamental biological processes. Moreover, they constitute unique models for investigations on how biomolecules are stabilized when subjected to extreme conditions. In addition, extremophiles offer a multitude of potential applications in various fields. Not only do many of them produce compounds of industrial interest, they also possess unique physiological properties.
Salt-Tolerant Alkaliphilic Actinomycetes Many alkaliphilic bacteria and few archaea live under the extreme range of pH, from pH 9 to 12 (Grant and Horikoshi, 1992). They rely on sophisticated transport mechanisms to maintain their intracellular pH near to neutrality by pumping or exerting protons. Among the *
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bacteria, a mycelial growth is the characteristic among the actinomycetes, a diverse group of Gram-positive bacteria. Actinomycetes have attracted greater due to their various natural products and specific mechanism of adapting extreme environments. While actinomycetes from normal habitats have focused considerable attention during the last many decades, exploration of such organisms from extreme habitats, particularly saline and alkaline, is a relatively new horizon. There are only few reports in literature about these microbes (Nyyssola et al., 2001; Al-Zarban et al., 2002a, 2002b; Hozzein et al., 2004; Kim et al., 2005; Starch et al., 2005, Montalavo et al., 2005). This has been mainly due to the difficulty associated with their isolation and maintenance under laboratory conditions. Recent culture independent studies have shown that marine environments contain a high diversity of actinobacterial species, if rediscovered by cultivation-based methods (Maldonado et al., 2005; Starch et al., 2005). Approximately 90% of the actinomycetes, cultured from saline and alkaline environments by using the unique techniques, were from the prospective new genera, a result indicative of its high selectivity. It is, therefore, relevant to pay further attention to extreme actinomycetes, as a possible way to discover novel taxa and, consequently, new secondary metabolites. Besides, they may also provide unique systems to investigate adaptive strategies and stability of macromolecules. The diversity displayed by them in terms of morphology, biochemical and molecular characters provides a greater insight into the microbial heterogeneity prevailing among them under extreme environments. Microbial natural products remain an important resource yet microorganisms from extreme environment have largely been overlooked in this regard. The recent discovery of novel primary and secondary metabolites from taxonomically unique populations of extremophilic actinomycetes suggests that these organisms add an important new dimension to microbial natural product research (Jensen et al., 2005). Continued efforts to characterize halo-tolerant alkaliphilic actinomycete diversity and how adaptations to the extreme environment affect metabolite production will create a better understanding of the potential utility of these organisms as a source of useful products for biotechnology. To date, the findings and research on salt-tolerant and alkaliphilic actinomycetes is largely based on phylogeny and only limited attempts have been made to explore their enzymatic potential and other biotechnological implications. They hold significance in the field of enzymology, pharmaceuticals, degradation of biomolecules, food technology and microbially enhanced oil recovery (Horikoshi, 1999; Basaglia et al., 1992; Abd-Allah, 2001; Kampfer et al., 2002).
Diversity of Alkaliphilic Actinomycetes The term “alkaliphile” is used for microorganisms that grow optimally or very well at pH above 9 but cannot grow or grow only slowly at the near-neutral pH value of 6.5. Alkaliphiles include prokaryotes, eukaryotes, and archaea. Many different taxa are represented among the alkaliphiles, and some of these have been proposed as new taxa (Horikoshi K., 1999). It is believed that the actinomycete complex of alkaline soils is dominated by many novel genera of actinomycetes including Streptomyces, which showed maximal radial rates of colony growth at pH 9-10. The research on actinomycetes surviving under extreme environments is still limited. However, alkaliphilic actinomycetes have gained
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considerable attention in recent years. There is tremendous diversity and novelty among the alkaliphilic actinomycetes present in alkaline habitats including soda lake and desert soils (Al-Zarban et al., 2002a,b; Li et al., 2003a,b, 2004, 2005, 2006a,b; Hozzein et al., 2004). A number of novel taxa have also been described by using polyphasic analysis and several molecular techniques.
Antimicrobial Potential of Halophilic and Alkaliphilic Actinomycetes Actinomycetes, soil bacteria with fungal-like filaments, have long been tapped by pharmaceutical researchers as a source of novel antibiotics, actinomycin and streptomycin for instance. Progress has been made to isolate novel actinomycetes from samples collected at different marine environments and alkaline habitats. These actinomycetes produce different types of new secondary metabolites. Many of these metabolites possess biological activities and have the potential to be developed as therapeutic agents. Marine actinomycetes are a prolific but underexploited source for the discovery of novel secondary metabolites including some highly biologically active substances (Lam, 2006). The phylogenetic analysis of the ketosynthase (KS) gene sequences of halophilic actinomycete Salinispora indicated that such strains might synthesize rifamycin-like compounds (Kim et al., 2006). Liquid chromatography- tandem mass spectrometry (LC/MS/MS) analysis demonstrated that Salinispora does produce compounds of the rifamycin class, including rifamycin B and rifamycin SV. This is the first recorded source of rifamycins from halophilic actinomycete. Similarly, William Fenical and his colleagues have discovered a new source of potential drugs - actinomycetes living in tropical and subtropical ocean sediments. They obtained the compound “salinosporamide A” from the Salinospora species of halophilic actinomycetes. The compound is believed to be a potent anticancer drug as it displayed strong and selective cytotoxic activity against cancer cells (Bradley, 2003). Dietera et al., (2003) characterized pyrocoll, an antibiotic, antiparasitic and antitumor compound produced by a novel alkaliphilic Streptomyces strain. Thus, further advances in the discovery and characterization of novel antimicrobial compounds from extremophilic actinomycetes can be considered as an aided feature in this research field.
The Enzymatic Potential of Salt-Tolerant Alkaliphilic Actinomycetes Because of increasing industrial demands for biocatalysts that can cope with industrial process conditions, considerable efforts have been devoted to the search for such enzymes. Despite the fact that to date more than 3000 different enzymes have been identified and many have been used in biotechnological and industrial applications, the available enzymatic array is still not sufficient to meet the ever inceasing demand. During the last few years, some extracellular enzymes from halophilic and alkaliphilic bacteria have been studied. However,
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little is known about the same with regards to extremophilic actinomycetes in general and salt-tolerant alkaliphilic actinomycetes in particular. They have been investigated to secrete a range of extracellular enzymes such as alkaline protease, amylase, lipase and cellulase.
Alkaline Proteases
Alkaline proteases constitute a very large and complex group of enzymes; with both nutritional and regulatory roles in nature. Proteases are among the commercially most viable enzymes and exploration of further novel microbial sources of this enzyme has enthused scientific community during the last several years. Proteases are among the most important class of industrial enzymes, which constitute >65% of the total industrial applications such as food additives, leather (dehairing agents), enhanced oil recovery, hyper saline waste treatment and in detergent formulation. Several haloalkaliphilic bacteria have been looked for there ability to secrete alkaline protease (Johanvesly et al., 2001, Kanekar et al., 2002; Nascimento and Martins, 2004; Patel et al, 2006); however, similar citation with reference to halophilic and alkaliphilic actinomycetes are relatively scares in the literature. Tsuchiya and coworkers (1997) carried out the detection and preliminary enzymatic characterization of intracellular alkaline protease from alkaliphilic Thermoactinomycets sp. HS682. Studies concerned with production and purification of alkaline protease were carried out from Streptomyces clavuligerus (Moreira et al., 2001). More recently, extracellular serine proteases secreted by an alkaliphilic actinomycete have been reported (Mehta et al., 2006).
Alkaline Amylases
Different types of amylases are found through out the animal, plant and microbial kingdom. α -Amylases, particularly of microbial origin are widely used in various industrial processes such as starch liquefaction, pulp processes and detergent formulation. Ammar and colleagues (2002) explained new action pattern of a maltose-forming alpha-amylase from alkaliphilic Streptomyces sp. and its possible application in bakery.
Alkaline Cellulases
Cellulose, the most abundant component of plant biomass, is found in nature almost exclusively in plant cell walls, although it is produced by some animals (e.g., tunicates) and a few bacteria. Fungi have traditionally been the main cellulase-producing microorganisms (Rajendran et al., 1994); though a few bacteria and actinomycetes have also been known to yield cellulase activity (Dasilva et al., 1993; Singh et al., 2004). The potential of cellulases has been revealed in various industrial processes, including food, textiles and laundry, pulp and paper, and agriculture as well as in research and development. Commercially available cellulases display optimum activity over a pH range from 4 to 6. No enzyme with an alkaline optimum pH for activity (pH 10 or higher) was reported before the rediscovery of
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alkaliphiles. Damude et al. (1993) studied a semi-alkaline cellulase produced by alkaliphilic Streptomyces strain KSM-9. Dasilva et al. (1993) reported cellulase from alkaliphilic Streptomyces sp. strain S36-2 which was active in the range of pH 8-9. Cellulase production has also been reported in an alkaliphilic Streptomyces strain S36-2. Van and coworkers described the cloning and expression of an endocellulase gene from a novel alkaliphilic Streptomycete isolated from an East African soda lake (2001).
Alkaline Lipases
Watanabe et al. (1977) conducted an extensive screening for alkaline lipase-producing microorganisms from alkaline soil and other habitats. Till date, lipolytic enzymes have been studied extensively from Pseudomonas and related oil degrading bacteria (Deb et al., 2006), while actinomycetes have been overlooked in this context. However, some recent investigations on the lipases from these groups of organisms are evident in the literature. Streptomyces strain SAP 1089 (Jain et al., 2003) produced alkaline lipases which were active and stable at pH 9-10 and highly activated by 0.14 mol/L CaCl2. Lescic and coworkers (2004) carried out the structural characterization of extracellular alkaline lipase from Streptomyces rimosus and assigned their disulfide bridge pattern by mass spectrometry. Similarly, a novel extracellular alkaline lipase from Streptomyces rimosus R6-554W hasa been purified and biochemically characterized with specific reference to cloning, sequencing, and high-level expression of its gene. This lipase showed no overall amino acid sequence similarity to other lipases in the databases. Besides these enzymes, few alkaliphilic actinomycetes have also been explored for their potential to secrete xylanases (Mansour et al., 2003) and chitinases (Gupta et al., 2005). Garg and coworkers (1996, 1998) reported a biobleaching effect of Streptomyces thermoviolaceus xylanase on birch wood kraft pulp. The enzyme was active at pH 9 and 65ºC. Extracellular xylanase from thermophilic Streptomyces sp.K37 was purified by ultrafiltration and cation exchange chromatography followed by gel filtration chromatography (Mansour et al., 2003). Tsujibo et al. (1992) isolated two types of chitinases from the alkaliphilic actinomycete, Nocardiopsis albus subsp. prasina OPC-131. The optimum pH of chitinase A was 8 and that of chitinase B, 7. The characterization of these Chitinase genes has also been carried out (Tsujibo et al., 2003).
Conclusion The member of halotolerant alkaliphilic actinomycetes are currently in culture is limited. The challenge today is to isolate, purify and cultivate microorganisms that have been so far remained “uncultivated”. The challenge will be to taxonomically classify them using both classical and molecular methods and exploit their genetic potential to yield novel bioactive molecules. So, it is of great value to discover new organisms from unexplored regions by developing suitable enrichment techniques and to study their unique enzymatic properties. In addition, isolation of these microorganisms will enlarge our knowledge of the “unseen
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majority” and will extend our view on microbial biodiversity beyond the presently described physiochemical boundaries for microbial growth. The application of halotolerant alkaliphilic actinomycetes and their metabolites in industrial processes has opened a new era in biotechnology. Further, the studies on alkaliphiles have led to the discovery of many types of enzymes that exhibit interesting properties and may generate a pool of new applications. The advances in the application of alkaliphilic- or alkalitolerant-based biomolecules during the past 20 years are in the center stage due to the introduction of proteolytic enzymes in the detergent industry. Industrial applications of alkaliphiles have been investigated and some enzymes have been commercialized. Of the enzymes now available to industry; proteases, cellulases, lipases and pullulanases are by far the most widely employed. Our own studies have indicated the wide occurrence of hydrolytic enzymes; such as alkaline protease, amylase and cellulase in salt-tolerant alkaliphilic actinomycetes isolated from vast coastal line and saline habitats along the Saurashtra Coast (Western India). Besides their diversity, the enzymatic potential has been clearly displayed. The combination of extreme conditions under which the biocatalyst are able to function would be of particular interest for developing novel processes and investigating structure and function relationship. The cloning and expression systems can also be developed for the production of enzyme in large quantity and it may help improving and tailoring the biocatalytic properties by protein engineering and random mutagenesis. Recent advances in genomics and related experimental technologies such as gene array and proteomics can be helpful in changing the approaches to the development and use of extremophilic actinomycetes as hosts for the production of both homologous and heterologous gene products.
Acknowledgemnet The work highlighted from our own group was sponsored by Saurashtra University and UGC, Government of India.
References Abd-Allah, E.F. (2001).Streptomyces pliicatus as a model bioconrol agent. Folia Microbiol., 46(4), 309-14. Al-Zarban, S.S., Abbas, I., Al-Musallam, A.A., Steiner, U., Stackebrandt, E. & Kroppenstedt, RM. (2002a). Nocardiopsis halotolerans sp. nov., isolated from salt mars soil Kuwait. Int J Syst Evol Microbiol., 52 (2), 525-529. Al-Zarban, S.S., Al-Musallam, A.A., Abbas, I., Stackebrandt, E. & Kroppenstedt, RM (2002b). Saccharomonospora halophila sp. nov., a novel halophilic actinomycete isolated from marsh soil in Kuwait. Int J Syst Evol Microbiol., 52(2), 555-558. Ammar, Y.B., Matsubara, T., Ito, K., Lizuka, M., Limpaseni, T., Pongasawasdi, P. & Minamiura, N. (2002). New action pattern of a maltose-forming alpha-amylase from
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Streptomyces sp. and its possible application in bakery. J Biochem Mol Biol., 35 (6), 568-75. Basaglia, M.G., Concheri, S., Cardinial, M.B., Pasti, G. & Nuti, M.P. (1992). Enhanced degradation of ammonium pre-treated wheat straw by lignocellulolytic Streptomyces spp. Can J of Microbiol., 38(10), 1022-1025. Bradley, D. (2003). Marine bugs make drugs. Angew Chem Int., 42(3), 355-357. Damude, H.G., Gilkes, N.R., Kilburn, D.G., Mille, R.C., Antony, R. & Warren, J. (1993). Endoglucanase CasA from alkalophilic Streptomyces strain KSM-9 is a typical member of family B of β-1, 4-glucanases. Gene., 123, 105–107. Dasilva, R., Yim, D.K., Asquieri, E.R. & Park, Y.K. (1993). Production of microbial alkaline cellulase and studies of their characteristics. Rev Microbiol., 24, 269-274. Deb, C., Daniel, J., Sirakova, T.D., Abomoelak, B., Dubey, V.S. & Kolattukudy, P.E. (2006). A novel lipase belonging to the hormone-sensitive lipase family induced under starvation to utilize stored triacylglycerol in Mycobacterium tuberculosis. J Biol Chem., 281(7), 3866-75. Dietera, A., Hamm, A., Fiedler, H.P., Goodfellow, M., Muller, W.E., Brun, R. & Bringmann, G. (2003). Pyrocoll, an antibiotic, antiparasitic and antitumor compound produced by a novel alkaliphilic Streptomyces strain. J Antibiot., 56, 639-46. Grant, W.D. & Horikoshi, K. (1992). Alkaliphiles: ecology and biotechnological applications. In Herbert, R.A. & Sharp R.J. (Eds), Molecular Biology and Biotechnology of extremophiles (pp.143-162). London: Blackie & son. Horikoshi, K. (1991). General view of alkaliphiles and thermophiles. In K.Horikoshi and W.D.Grant (Eds.), super bugs: Mocro-organisms in Extreme Environments (pp. 3-13). Springer Verlag Erlin. Horikoshi, K. (1999). Alkaliphiles: some applications of their products for biotechnology. Microbiol Mol Biol Rev., 63 (4),735-743. Hozzein, W.N., Li, W.J. , Ibrahim, A.M., Hammouda, O., Mousa, A.S., Xu, L.H. & Jiang, C.L. (2004). Nocardiopsis alkaliphila sp. nov., a novel alkaliphilic actinomycete isolated from desert soil in Egypt. Int J Syst Evol Microbiol., 54, 247-252. Jensen, P.R., Mincer, T.J., Williams, P.G. & Fenical, W. (2005). Marine actinomycete diversity and natural product discovery. Antonie Van Leeuwenhoek, 87(1), 43-48. Johnvesly, B. & Naik, G.R. (2001). Studies on production of thermostable alkaline protease from thermophilic and alkaliphilic Bacillus sp. JB-99 in a chemically defined medium. Process Biochem., 37, 139-144. Kampfer P., Busse HJ. & Rainey FA. (2002). Nocardiopsis compostus sp. nov., from the atmosphere of a composting facility. Int J Syst Evol Microbiol., 52(2), 621-627. Kanekar, P.P., Nilegaonkar, S.S. Sarnaik, S.S. & Kelkar, A.S. (2002). Optimization of protease activity of alkaliphilic bacteria isolated from an alkaline lake in India. BioresTechnol.,85(1), 87-93. Kim, T.K., Hewavitharana, A.K., Shaw, P.N. & Fuerst, J.A. (2006). Discovery of a new source of rifamycin antibiotics in marine sponge actinobacteria by phylogenetic prediction. Appl Environ Microbiol.,72(3), 2118-25.
Salt-tolerant Alkaliphilic Actinomycetes and their Biocatalytic Potential
227
Kim,T.K., Garson, M.J. & Fuerst, J.A. (2005). Marine actinomycetes related to the “Salinospora” group from the Great Barrier Reef sponge Pseudocaratina clavata. Environ Microbiol., 7(4), 509-18. Lam, K.S. Discovery of novel metabolites from marine actinomycetes. Curr Opin Microbiol. 2006, 9(3): 245-51. Lescic, I., Zehl, M., Muller, R., Vukelic, B., Abramic, M., Pigac, J., Allmaier, G. & KojicProdic, B. (2004). Structural characterization of extracellular lipase from Streptomyces rimosus: assignment of disulfide bridge pattern by mass spectrometry. Biol Chem , 385(12), 1147-56. Li, W.J., Chen, H.H., Zhang, Y.Q., Schumann, P., Xu, L.H. & Jiang, C.L. (2004). Nesterenkonia halotolerans sp. nov. and Nesterenkonia xinjiangensis sp. nov., actinobacteria from saline soils in the west of China. Int J Syst Evol Microbiol., 54, 837841. Li, W.J., Kroppenstedt, R.M., Wang, D., Tang, S.K., Lee, J.C., Park, D.J., Kim, C.J., Xu, L.H. & Jiang, C.L. (2006a). Five novel species of the genus Nocardiopsis isolated from hypersaline soils and emended description of Nocardiopsis salina Li et al. 2004. Int J Syst Evol Microbiol., 56(5), 1089-96. Li, W.J., Tang, S.K., Stackebrandt, E., Kroppenstedt, R.M., Schumann P., Xu, L.H. & Jiang, C.L. (2003b). Saccharomonospora paurometabolica sp. nov., a moderately haophilic actinomycete isolated from soil in China. Int J Syst Evol Microbiol., 53(5), 1591-4. Li, W.J., Xu, P., Zhang, L.P., Tang, S.K., Cui, X.L., Mao, P.H., Xu ,L.H., Schumann, P., Stackebrandt, E. & Jiang, C.L. (2003a).Streptomonospora alba sp.nov., a novel halophilic actinomycete, and emended description of the genus Streptomonospora Cui et al., 2001. Int J Syst Evol MIcrobiol., 35, 1421-1425. Li, W.J., Zhang, Y.G, Zhang, Y.Q., Tang, S.K., Xu, P., Xu, L.H. & Jiang, C.L. (2005). Streptomyces sodiiphilus sp. nov., a novel alkaliphilic actinomycete. Int J Syst Evol Microbiol., 55, 1329-1333. Li, W.J., Zhang, Y.Q., Schumann, P., Chen, H.H., Hozzein, W.N., Tian, X.P., XU, L.H. & Jiang, C.L. (2006b). Kocuria aegyptia sp. nov., a novel actinobacteria isolated from a saline, alkaline desert soil in Egypt. Int J Syst Evol Microbiol., 56 (4), 733-7. Maldonado, L.A., Starch, J.E., Pathom-aree, W., Ward, A.C., Bull, A.T. & Goodfellow, M. (2005). Diversity of cultivabe actinobacterial in geographically widespread marine sediments. Antonie Van Leeuwenhoek, 87(1), 11-18. Mehta, V.J., Thumar, J.T. & Singh, S.P. (2006). Production of alkaline protease from an alkaliphilic actinomycete. Bioresour Technol., 97(14), 1650-4. Montalavo, N.F., Mohamed, N.M., Enticknap, J.J. & Hill, R.T. (2005). Novel actinobacterial from marine sponges. Antonie Van Leeuwenhoek, 87(1), 29-36. Moreira, K.A., Cavalcanti, M.T.., Duarte. H.S., Tambourgi, E.B., Magalhães de Melo, E.H., Silva, V.L., Porto, A.L. & Filho, J.L. (2001). Partial characterization of proteases from streptomyces clavuligerus using an inexpensive medium. Braz J Microbiol., 32, 623-629. Nascimento, W.C. & Martins, M.L. (2004). Production and properties of an extracellular protease from thermophilic Bacillus sp. Braz J Microbiol., 35, 1-2. Nyyssola, A. & Leisola, M. (2001). Actinopolyspora halophila has two separate pathways for betaine synthesis. Arch Microbiol., 176(4), 294-300.
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Patel, R.K., Dodia, M.S., Joshi, R.H. & Singh, S.P. (2006). Purification and characterization of alkaline protease from a newly isolated haloalkaliphilic Bacillus sp. Process Biochem., 41, 2002-04. Rajendran, A., Gunashekaran, P. & Lakshmanan, M. (1994). Cellulase activity of Humicola fuscoatra. Ind J Microbiol., 34(3), 289-295. Singh, J., Batra, N., & Sobti, C. (2004a). Purification and characterization of alkaline cellulase produced by a novel isolate, Bacillus sphaericus JS1. J Ind Microbiol Biotechnol., published on line. Starch, J.E. & Bull, A.T. (2005). Estimating and comparing the diversity of marine actinobacteria. Antonie Van Leeuwenhoek, 87(1), 3-9. Tsuchiya, K., Ikeda, I., Tsuchiya, T. & Kimura, T. (1997). Cloning and expression of an intracellular alkaline protease from alkaliphilic Thermoactinomycets sp.HS682. Biosci Biotechnol Biochem., 61(2), 298-303. Van Solingen, P., Meijer, D., Van der Kleij, W.A., Barnett, C., Bolle, R., Power, S.D. & Jones, B.E. (2001). Cloning and expression of an endocellulase gene from a novel Streptomycete isolated from an East African soda lake. Extremophiles., 5(5),333-41. Watanabe, N., Ota, Y., Minoda, Y. & Yamada, K. (1997). Isolation and identification of alkaline lipase-producing microorganisms, culture conditions and some properties of crude enzymes. Agric Biol Chem., 41, 1353–1358.
In: Biocatalysis Research Progress Editors: F. H. Romano, A. Russo
ISBN: 978-1-60456-619-2 © 2008 Nova Science Publishers, Inc.
Chapter IX
Accelerating Whole-cell Biocatalysis by Cellular Membrane Engineering Ye Ni1,*, Rachel R. Chen2,* 1
Laboratory of Biocatalysis, School of Biotechnology, Jiangnan University, The Key Laboratory of Industrial Biotechnology, Ministry of Education, Wuxi 214122, P.R. China 2 School of Chemical & Biomolecular Engineering, Georgia Institute of Technology Atlanta, Georgia 30332-0100, USA
Abstract Whole-cell biocatalysts are preferred in biocatalysis applications involving cofactors and/or multiple enzymes. However, cell envelope often represents a formidable permeability barrier, limiting the rate of entry of substrate. As a result, reactions catalyzed by whole-cells are reportedly orders of magnitude slower than those of by their free enzyme counterparts. This chapter reviews recent molecular engineering efforts addressing this critical issue. Initial studies were carried out with E. coli strains carrying mutations in the outer membrane structures. The effects of these mutations were investigated by interrogating the mutant cells with substrates differing substantially in size and hydrophobicity. The reduction of outer membrane permeability barrier by these mutations led to significant accelerations in reaction rates of all whole-cell catalyzed reactions investigated. In the case of the tetrapeptide, a substrate for subtilisin, a single gene mutation in lipopolysaccharide (LPS) synthesis can render the outer membrane completely permeable to substrate, reaching a barrier-less condition that maximizes the reaction rate while retaining all the benefits of whole-cell catalysts. For reaction rates of toluene
* *
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Ye Ni and Rachel R. Chen dioxygenase (TDO)-catalyzed reactions, an increase of up to 6-fold was observed with lipoprotein mutant for three small, hydrophobic substrates tested. Mutations in either LPS or lipoprotein are effective for accelerating reactions with UDP-glucose, a hydrophilic molecule with Mw over 600 Da, resulting in a striking acceleration (up to 14-fold) of reaction rate. The magnitude of reaction rate acceleration was found to be dependent upon the substrate concentrations, the enzyme expression level, and the nature of the mutations. In addition, the mutations were demonstrated to be far more superior to common permeabilization procedures such as freeze-thaw and EDTA treatments. To understand the mechanism of the permeability enhancement in the lipoprotein mutant, the lpp region was sequenced. The results revealed that Braun’s lipoprotein was absent in the transposon mutant cells due to multiple stop codons within the 59-bp insertion, suggesting that the absence, rather than the alteration of Lpp, is responsible for the observed change in permeability. More importantly, the sequencing result suggests that lpp deletion could become a general permeabilization method. Subsequent studies were carried out by generating lpp deletion mutants from strains with different genetic background. It was indeed shown that lpp deletion generates a useful phenotype, not only effective in enhancing substrate permeability, but also in reactions limited by product permeability, as demonstrated in L-carnitine synthesis. Importantly, the deletion has no significant effect on cell growth, metabolism and recombinant protein expression. Therefore, lpp deletion phenotype could be applied as a generally applicable method to enhance the outer membrane permeability of various E. coli strains, and possibly other Gram-negative bacteria with lpp homologs.
Introduction Compared to isolated enzymes, whole-cell biocatalysis has a number of attributes that are particularly attractive for large-scale applications. First, the use of whole cells is more economical as it eliminates the need for tedious, expensive protein isolation and purification. Second, whole cell biocatalysis allows cascades of enzymatic reactions that involve multiple enzymes, cofactors, and substrates while the same reaction might be too complicated to perform using isolated enzymes. Moreover, enzymes are generally more stable when protected by cell envelopes [1, 2]. However, the reactions catalyzed by whole cells are generally 10– to 100–fold slower than rates observed with free enzyme reactions. The difference is attributed to the mass transport barrier imposed by cellular envelopes. For example, isonovalal synthesis from αpinene oxide catalyzed by Pseudomonas rhodesiae whole cells was increased by 60–fold after permeabilizing the cells [3]. In another recent study, mass transfer across a recombinant E. coli cell membrane was shown to cause a ten-fold reduction in the reaction rate of BaeyerVilliger oxidations of biocyclo[3.2.0] [4]. In a more dramatic case, intact Pseudomonas pseudoalcaligenes cells showed no activity to maleate (substrate) in the process of D-malate production; the cell membrane completely blocked the entry of the substrate [5]. Numerous other studies reported similar mass transfer limitations in whole-cell biocatalysis [6, 7, 8, 9]. This common mass transport problem has thus far limited the commercial application of whole-cell biocatalysis. Until recently, the only viable strategy to address the issue was
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permeabilization [10]. General permeabilizing methods involve chemical treatments with detergents, solvents, and chelators, or physical treatments such as extreme temperature fluctuations. All these methods are highly empirical, and are generally undesirable for large scale applications. For example, permeabilizing cells with 1% toluene, a commonly used procedure, requires five steps [8]. Permeabilization treatments inevitably create additional steps in manufacturing processes, complicate downstream processing especially when a surfactant is used, and incur unintended damage to cells. Cell damage is particularly problematic as it disrupts the normal membrane functions and intracellular metabolism needed for efficient cofactor regeneration. With the advent of recombinant DNA technology, molecular engineering strategies emerged in recent years to address the permeability issue in whole-cell bioprocesses. This chapter reviews the progress of using molecular engineering tools to reduce the permeability barrier and thereby enhancing biocatalysis rates. The effects of two E. coli outer-membrane mutations (lipopolysaccharide and Braun’s lipoprotein) on the permeability of substrates/products were studied. The molecular engineering approach was compared with permeabilization methods such as EDTA treatment and freeze-thaw. The genetic sequencing of the transposon mutant of Braun’s lipoprotein was carried out. The result showed that the Lpp was not expressed due to multiple stop codons within the insertion, prompting further investigation of the possibility of establishing a general permeabilization method by simple single gene deletion.
Molecular Basis of Permeability Limitation Microbial cell envelope, the first barrier to the noxious agents in the environment, has evolved to take up nutrients and exclude toxic chemicals. In Gram-negative bacteria cells, the outer-membrane of cell envelope (Figure 1A) is a distinct bilayered structure. It is chemically different from usual biological membranes and has the ability to resist damaging chemicals. The outer leaflet of the outer membrane, or lipopolysaccharide (LPS), is a highly ordered quasi-crystalline structure with very low fluidity and very low permeability to hydrophobic molecules. The LPS consists of lipophilic lipid A, an oligosaccharide core, and a long polysaccharide chain comprising up to 40 sugars (called an O-specific chain) (Figure 1B). The hydrophilic polysaccharide component is responsible for the exclusion of hydrophobic molecules and the hydrophobic lipid A of LPS limits the entry of hydrophilic compounds. The outer membrane of Gram-negative bacteria excludes hydrophobic compounds, regardless their size. For large hydrophilic molecules (>600–700 Daltons), dedicated protein-based mechanisms are needed for their entry whereas smaller hydrophilic molecules rely on passive diffusion through porin channels of the outer membrane. However, even with porins, diffusion may be slower than desired [9]. For biotechnology applications such as whole-cells biocatalysis, substrate/product is often synthetic, or hydrophobic and/or relatively big molecules. As a result, the import/export process is often the rate-limiting step of a biocatalysis reaction, and obtaining rapid permeation through outer membranes is a common and formidable obstacle to the commercialization of whole-cell biocatalysis applications.
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(A)
(B) Figure 1. (A): Gram-negative envelope. (B): Lipopolysaccharide structure. (Adopted from Prescott et al., 2002 [11]).
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Whole-cell Reactions with β-lactamase and Subtilisin The effects of the two outer membrane mutations were investigated with β-lactamase and subtilisin using their respective substrates, nitrocefin and N-succinyl-Ala-Ala-Pro-Phe-pnitroanilide (referred as tetrapeptide in the text). The two mutants are E. coli mutant (SM101) [12 13 14 15 ] , , , , carrying a mutation in the synthesis of lipopolysaccarides, and E609L [16, 1], carrying an insertion in the lpp gene (encoding Braun’s lipoprotein). Their respective parental strains (SM105 and E609) were used as controls. In order to focus our study on outer membrane permeability, we used two plasmids that directed the expression of two enzymes of interest into the periplasmic space. Plasmid pBR322 carries a gene coding for β-lactamase along with its signal sequence that allows secretion of the expressed enzyme to the periplasmic space [17]. Plasmid pGES201 allows periplasmic expression of subtilisin, a serine protease [18]. By measuring the whole-cell activities of the β-lactamase and subtilisin, we were able to quantify the effects of the mutations on reaction rates thereby the permeability of their respective substrates, nitrocefin (MW=516.5, hydrophilic) and N-succinyl-Ala-Ala-Pro-Phep-nitroanilide (a tetrapeptide, MW=624.6, hydrophobic). For nitrocefin, similar β-lactamase activities of cell-free extract for all four constructs indicate neither mutation significantly affects the ability of the cells to synthesize the recombinant enzymes. Whole cells had only 5%–to–20% of the cell-free extract activity, indicating permeability limitation for the substrate. LPS mutation increased the activity of the whole cells by about 50% while lipoprotein mutation increased the activity by 3.8–fold (Figure 2A). For reaction with tetrapeptide (AAPF-pNA), both mutants had lower expression of the enzyme than the wild types (15% and 25% lower, respectively). Despite the lower expressions, mutant whole-cells exhibited activities either similar to (E609 pair) or significantly higher (SM pair) than the activities obtained by their respective parental strains, indicating increased permeability by either mutation. Mutation of LPS increased the wholecell activity to 95% of the cell-free extract level (Figure 2B). This result is significant as it implies that the mutation rendered the whole-cells completely permeable to the tetrapeptide and eliminated the mass transport resistance, a membrane-less or barrier-less condition that maximizes reaction rates.
Whole-cell Reactions with Toluene Dioxygenase Toluene dioxygenase (TDO) [19] is a multi-component enzyme system that catalyzes the dioxygenation of toluene to toluene cis-dihydrodiol. TDO was known to be active toward toluene and its related structures including ethylbenzene and 2-indanone. Taking advantage [1]
Miller KW, Schamber R, Chen Y, Ray B. (1998). Production of active chimeric pediocin AcH in Escherichia coli in the absence of processing and secretion genes from the Pediococcus pap operon. Appl Environ Microbiol, 64:14-20
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of its broad substrate range, the mutation effect on permeability of the three small hydrophobic substrates (toluene, ethylbenzene and 2-indanone) was studied.
A 25 Percentage Activity (%)
Nitrocefin 20
SM101/pBR322 SM105/pBR322
15
E609L/pBR322 E609/pBR322
10 5 0
LPS mutation
Lipoprotein mutation
B Percentage Activity (%)
100 Tetrapeptide 80 SM101/pGES201
60
SM105/pGES201 E609L/pGES201
40
E609/pGES201
20 0 LPS mutation
Lipoprotein mutation
Figure 2. Effects of LPS (A) and lipoprotein (B) mutations on the permeability of nitrocefin and tetrapeptide. Percentage activities are calculated by dividing whole cell activities by their respective cell-extract activities [20] .
Figure 3 shows a comparison of initial reaction rates between the mutant and its wildtype control for toluene substrate. Over the range of toluene concentration tested, the initial rate ratios of mutant to parent strains varied over a range of 1.5 (at 0.1 mM) to 3.9–fold (at 5 mM). Similar rate enhancement was observed for other two substrates (data not shown). Possible substrate and/or product inhibitions was observed for both parent and mutant strains. Interestingly, the mutant strain seemed to fare better in terms of inhibition and only a slight
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decrease of reaction above substrate concentration of 4 mM was evident (Figure 3). For ethylbenzene, the initial velocity-substrate concentration profile was similar to that of toluene except that there was a sharp decline of enzyme activities at high substrate concentrations for both strains. For 2-indanone, the mutant strain had a substantially higher reaction rate than that of the parent over the entire range of concentrations examined. No inhibition was evident. Although some of the components of TDO are inner membrane bound and some reside intracellularly, toluene, ethylbenzene, and 2-indanone are sufficiently hydrophobic and their passage through inner membrane is not expected to be rate-limiting [9]. The increased initial reaction rate is therefore likely through improved outer membrane permeability to substrates due to the mutation. Additional time course studies were carried out to further evaluate the potential of the lipoprotein mutant as whole-cell biocatalyst in bioremediation and green chemistry applications. The results show that the mutant demonstrated approximately 75% conversion ratio, whereas only about 30% for the parent strain. The observed difference in conversion and product concentration was consistent with the observed faster initial velocity (Figure 3) for the mutant. In all concentrations tested, the mutant degraded more toluene and achieved higher product concentrations. At an initial concentration of 1 mM toluene, for example, nearly 100% toluene was degraded by the mutant while the parent could only degrade about 50%. Similar results were obtained with other two substrates. EDTA, an effective membrane permeabilizer, has long been used to treat bacterial cell membranes to increase permeability [21]. It functions through chelation of metal ions such as Ca++ and Mg++, which cross-link lipopolysaccharides, consequently adding to the rigidity of the outer membrane. Since the LPS layer is suspected to be the main permeability barrier to hydrophobic molecules such as toluene, removal of these metal ions is expected to weaken the intra-molecular interaction of LPO and loosen up the outer membrane layer, thus allowing passage of hydrophobic molecules through the outer membrane layer at a higher rate. Our studies showed that EDTA treatment increased reaction rate with toluene for about 23% (±5% SD) for the parent strain E609 under optimal conditions (0.5 mM EDTA, 15 min) (Figure 4). This compares with the lipoprotein mutation that increased the reaction rate more than two fold (Figure 3), indicating the mutation in envelope structure could be far superior to chemical treatment. Increasing EDTA concentration higher than 0.5 mM caused a decrease of TDO activities, probably due to the removal of the required metal cofactors for TDO (Fe++) and the destabilizing of the inner membrane. This result further illustrates that the chemical treatment causes undesired damage. Interestingly, no increase in reaction rate by EDTA was observed for the mutant strain. The treatment, in fact, caused a sharp decrease of TDO activities (Figure 4). This is probably due to the defect in membrane structure caused by the lipoprotein mutation that sensitizes the cells for EDTA treatment [16].
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Activity (mUnits/1.0 OD600 cell)
6 5 4 3 2 1 0 0
1
2
3
4
5
Conc. of Toluene (mM) Figure 3. Effect of toluene concentration on the initial degradation rate by whole- cell E. coli E609L/pDTG601 (▲) and E609/pDTG601 (■). Error bars represent standard deviations from the mean. All data are measured in at least three separate experiments [22].
6 mUnits/1.0 OD600 cell
Control 5
0.1 mM EDTA 0.5 mM EDTA
4 3 2 1 0 E609L
E609
Figure 4. Effect of EDTA on the permeability of E609L/pDTG601 and E609/pDTG601. Error bars represent standard deviations from the means. All data are measured in at least three separate experiments [22].
Whole-cell Reactions with UDP– Glucose Dehydrogenase Permeability study with E. coli cells that express recombinant UDP-glucose dehydrogenase, was also performed. UDP-glucose is a relatively large molecule (MW=610), close to the maximum size allowable through porins9. In addition, since it is hydrophilic, it provides a useful contrast to our earlier studies with small hydrophobic molecules.
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Activity (mUnits/1.0 ml 1.0OD600 Cell)
60 50 40 30 20 10 0 0
10
20
30
40
50
Conc. of UDP-Glc (mM) Figure 5. The effect of substrate concentrations on the initial rate of whole-cell E. coli SM101/pQEkfiD (S), SM105/pQEkfiD (U), E609L/pQEkfiD () and E609/pQEkfiD (
). 2 mM NAD+ was used in all reactions. Error bars represent standard deviations from the mean. All data are measured in at least three separate experiments [23].
The permeability of the membranes was evaluated by measuring the reaction rate catalyzed by an overexpressed enzyme, UDP-glucose dehydrogenase. Figure 5 shows a comparison of whole-cell activities between the mutants and the respective controls. Wholecells carrying either type of mutation exhibited a much higher activity than their respective parental strains, increasing reaction rate by a factor of 2.0–to–14, depending on the substrate concentrations used. Under all conditions, there were no significant differences in the expression levels of the enzyme between the mutant and the control as judged by both SDSPAGE and the activity measurement in the cell extracts. Therefore, the differences observed in the reaction rate can be attributed to the altered rate of substrate uptake due to the mutations. When whole-cell activities are compared to their cell extract activities, striking differences are evident (Table 1). At a relatively low substrate concentration (5 mM), the whole-cell activities for the parental strains only represent a small fraction of total activity inside the cells, about 4% for both E609 and for SM105, suggesting that severe permeability limitations imposed by the cell membranes. The mutations at the outer membrane structure dramatically increased the whole-cell activity, 9–fold for SM101 and 14–fold for E609L, bringing it to about 34% of the cell extract level for SM101 and 60% for E609L. At a higher substrate concentration (20 mM), the LPS mutant recovered about 73% of the activity, compared to about 19% with the parental strain. The lipoprotein mutant recovered nearly 100% of the cell extract activity whereas only about 10% activity was exhibited by the control strain. The results indicate that mutations at the outer membrane structures were effective in reducing the substrate permeability barrier in whole-cell catalysts. The
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lipoprotein mutation seemed to be particularly effective, capable of eliminating the permeability barrier under certain conditions. The effectiveness of outer membrane mutation in reducing substrate permeability limitations was compared to a common permeabilization procedure, freeze-thaw (FT), known to be effective in permeabilizing cell membranes for UDP-glucose [24]. Figure 6 shows that mutation of the LPS synthesis pathway is much more effective, increasing the whole-cell activity by about 9 times versus 2 times with FT. Similarly, the lipoprotein mutation led to an increase of nearly 14–fold, compared to about 5–fold with FT. Besides being more effective, the genetic engineering approach eliminates the need for additional processing steps, time, and equipment. Therefore, modification of the outer membrane is far superior to FT treatment. Table 1. A comparison of UDP-glucose dehydrogenase activity in whole-cell and cell extract of SM101/pQEkfiD, SM105/pQEkfiD, E609L/pQEkfiD and E609/pQEkfiD [23] Activitya (mUnits/1 ml 1.0OD600 cell) SM101 SM105 E609L E609 Cell extract
43.0 ± 2.2
39.9 ± 0.4
49.5 ± 1.6
51.7 ± 5.7
At 5 mM Substrate Concentration Whole cell
14.7 ± 1.7
1.6 ± 0.1
29.5 ± 2.6
2.1 ± 0.2
Whole cell/Cell extract (%)
34.2
4.0
59.6
4.1
At 20 mM Substrate Concentration Whole cell 31.4 ± 0.9 7.4 ± 0.7 48.8 ± 6.5 4.9 ± 1.1 Whole cell/Cell extract (%) 73.0 18.5 98.6 9.5 a All activities are measured in at least three separate experiments and expressed as means ± 1 standard deviation.
Activity Change (fold)
18 16
Freeze-Thaw
14
Mutation
12
14.0
9.2
10 8
4.9
6 4
2.2
2 0 SM105
E609
Figure 6. A comparison of membrane mutation and freeze-thaw method. 5 mM UDP-Glc and 5 mM NAD+ were used in all reactions. Error bars represent standard deviations from the mean. All data are measured in at least three separate experiments [23].
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The enzyme investigated, UDP-glucose dehydrogenase, was over-expressed in the cytoplasm. Our current system does not allow an assessment of the inner membrane permeability. As a matter of fact, our initial efforts in isolating the outer membrane permeability from inner membrane permeability (by expressing these enzymes in periplasmic space) resulted in either inactive enzymes or very low level expression, not conducive to the permeability study. Nevertheless, the significant rate acceleration achieved by mutations on the outer membrane structure suggests that the outer membrane permeability barrier for this molecule is significant and substantial reduction could be achieved through membrane engineering. If inner membrane permeability can be addressed, the maximum activity could be reached at an external substrate concentration lower than 20 mM.
Lpp Deletion as a Permeabilization Method The above permeability studies have demonstrated that Braun’s lipoprotein mutant E609L carrying a Tn10 insertion is particularly promising for whole-cell biocatalysis applications [20, 22, 23]. The remarkable merits of this lipoprotein mutation have prompted us to investigate this phenotype as a generally applicable permabilization method for biotechnological applications. However, E609L is not well-characterized. Braun’s lipoprotein exists in both free form and bound form with covalent linkage to the peptidoglycans [9]. The protein is subject to lipidation, which is necessary for the interaction of the cell envelope. Therefore, it is possible that the Tn10 insertion could lead to a structural change of the lipoprotein, with outcomes dependent upon the insertion location within the gene, and insertion sequences. It is also possible that the same mutation would lead to different outcomes, depending upon the genetic background of the strain. Previous permeability studies were carried out with E609 strain, a K-12 strain. This leads to a question whether useful phenotypes could be generated with other genetic background. In case that transposon excision occurs, the antibiotic marker could be incorporated into chromosome at a secondary location, which is the case with E609L. Thus, there is a chance that the observed effects of insertion could be caused by the secondary insertion. To ascertain that the permeability change and resulting phenotype is from the intended mutagenesis, and to understand the nature of the mutation, the lpp region of the insertional mutant was sequenced. Compared to its isogenic parent E609, the lpp gene in E609L contains a 59-bp insertion (grey shaded) 17 bp downstream from the start codon (Figure 7). While an insertion is expected, it is much shorter than expected (59 bp versus 9300 bp) and the antibiotic tetracycline resistance gene was not found within the insertion. Apparently, transposon excision occurred and the tetracycline resistant gene was inserted into an unknown location, which confers the strain tetracycline resistant. The 59 bp insertion contains a duplicate of 9 bp from the target gene (TACTAAACT), typical for a Tn10. It also contains an inverted repeat of 19 bp, characteristic of a Tn10. Further inspection of the inserted sequence revealed that the insertion contains two double-stop codons (in bold), indicating that the lpp gene would not be expressed at all in any case. Therefore, the absence of this major outer membrane lipoprotein, rather than the structural change, was responsible for the strikingly increased permeability observed earlier. Importantly, it suggests that the enhanced
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permeability could be obtained by a simple lpp deletion. Potentially, lpp deletion could be established as a generally applicable method for permeabilizing whole-cells for increased permeability thereby improved processes. E609 E609L
ACTCAATCTAGAGGGTATTAATAATGAAAGCTACTAAACT-------------------ACTCAATCTAGAGGGTATTAATAATGAAAGCTACTAAACTCTGATGAATCCCCTAATGAT ****************************************
E609 E609L
---------------------------------------GGTACTGGGCGCGGTAATCCT TTTGATAAAAATCATTAGGGGATTCATCAGTACTAAACTGGTACTGGGCGCGGTAATCCT *********************
E609 E609L
GGGTTCTACTCTGCTGGCAGGTTGCTCCAGCAACGCTAAAATCGATCAGCTGTCTTCTGA GGGTTCTACTCTGCTGGCAGGTTGCTCCAGCAACGCTAAAATCGATCAGCTGTCTTCTGA ************************************************************
E609 E609L
CGTTCAGACTCTGAACGCTAAAGTTGACCAGCTGAGCAACGACGTGAACGCAATGCGTTC CGTTCAGACTCTGAACGCTAAAGTTGACCAGCTGAGCAACGACGTGAACGCAATGCGTTC ************************************************************
E609 E609L
CGACGTTCAGGCTGCTAAAGATGACGCAGCTCGTGCTAACCAGCGTCTGGACAACATGGC CGACGTTCAGGCTGCTAAAGATGACGCAGCTCGTGCTAACCAGCGTCTGGACAACATGGC ************************************************************
E609 E609L
TACTAAATACCGCAAGTAATAGTACCTGTGAAGTGAAAAATGGCGCACATTGTGCGCCAT TACTAAATACCGCAAGTAATAGTACCTGTGAAGTGAAAAATGGCGCACATTGTGCGCCAT ************************************************************
Figure 7. Comparison of nucleotide sequences of lpp from E609L and E609Y. Start and stop codon are in bold. The 59-bp insertion part introduced by transposon is grey shaded [25].
Conc. of L-carnitine (mM)
35 30 25 20 15 10 5 0 0
4
8
12
16
20
24
Time (hr) Figure 8. Time course of L-carnitine production by whole-cell E. coli O44K74Y (
) and O44K74 (U), with 50 mM initial crotonobetaine and ~20 g wet cell/liter biomass. Error bars represent standard deviations from the mean. Data presented are average from three separate experiments [25].
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To investigate this possibility, several lpp deletion mutants were generated from various genetic backgrounds. The effect of the lpp deletion on the membrane permeability was first studied with a reaction catalyzed by UDP-glucose dehydrogenase with UDP-glucose as substrate, a system previously studied with E609L [23]. As expected, lpp deletion strain E609Y resulted in a 15–fold increase in the reaction rate, similar to what was observed before with E609L, indicating that the deletion generates a phenotype essentially the same as the transposon mutant. A hydrolysis reaction catalyzed by recombinant organophosphorus hydrolase (OPH) [26, 27, 28, 29]was also tested using E609Y to illustrate the versatility of lpp deletion as a permeabilization method. A 4-fold increase in the initial rate and 50% higher conversion in the deletion mutant can be attributed to the enhanced permeability through the outer membrane as the mutant and the wild type have similar expression level of the organophosphohydrolyase. In a study to illustrate how lpp deletion could enhance bioconversion rate in reactions where product permeability through the outer membrane is a limiting factor, L-carnitine [R-()-3-hydroxy-4-trimethylaminobutyrate] synthesis from crotonobetaine was investigated with the lpp deletion.mutant. A deletion mutant (O44K74Y) was generated from the high Lcarnitine producing strain, O44K74, [30, 31, 32, 33, 34]. Extensive earlier studies have established that the product permeability is important for a high yielding process [30, 31, 34]. Due to product permeability limitation, the yield was low (30–43%). Although chemical permeabilizers such as organic solvents, detergents, and polyethylenimine were effective in permeabilizing, they caused excessive damage to the cell envelopes [31]. Therefore, a permeabilization method that has minimal impact on cell physiology is ideal for this application. As shown in Figure 8, the initial rate of synthesis is high in the beginning of the reaction, and the difference in the initial rate between the mutant (O44K74Y) and the wild type (O44K74Y) is rather small. At 24 hours, the product concentrations reached were 28.0 mM and 21.6 mM, corresponding to a yield of 56% and 43% for the mutant and the wild type, respectively. This represents a 30% higher product yield for the mutant strain. Similar yield improvement (30-40% increase) was observed with higher substrate concentrations. The difference between the two strains was primarily observed toward the end of the reaction. This is reasonable because this reaction is product-permeability-limited. The enhanced product permeability due to lpp deletion and the effect on product yield were only observed when the product concentration was high, which occurred toward the end of the reaction.
Effect of Lipoprotein Mutation on Cell Growth and Recombinant Protein Expression One essential attribute for microbial whole-cell biocatalysts is their ability to grow and attain high cell density. Since the membrane function is critical to cell growth, any mutation aimed to increase the substrate permeability should be evaluated with respect to the effect on cell growth. Growth studies showed that the mutant cells grew as fast as the parent strain, even under stressful conditions such as carrying a medium-sized plasmid and overproducing a four-component recombinant enzyme system. It seems that the mutant cells were able to
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compensate the defect in membrane by some unknown mechanisms. The defect may affect the energetic state of the membrane, the proton gradient, for example. If this is true, it is conceivable that the mutant cells can compensate the loss of proton gradient by metabolizing more of an energy source, which would lead to a lower growth yield. Consistent with this explanation, we observed a slightly lower growth yield for the mutant in all three cases investigated (about 10−20% lower in cell concentrations). The mutant cells appear to be sensitive towards the end of the growth period, indicated by the loss of its ability to maintain cell biomass[22]. The exhaustion of an energy source, which mutant cells are relying on to maintain the membrane energetic state and compensate the defect, could be responsible for the observed behavior at the end of the cultivation. Overexpression of one or more recombinant enzymes is another essential attribute of the whole-cell catalyst. A full evaluation of the permeability barrier was not possible as attempts to remove the cell membrane barrier and reconstitution of TDO components in vitro failed to recover a majority of the enzyme activity. The SDS-PAGE gel showed that there is essentially no detectable difference in expression levels of TDO between the two strains for all components of the TDO system22. Therefore, the mutation in lipoprotein does not affect the expression of the multi-component TDO system.
Conclusion We demonstrate here that molecular engineering approach is effective in addressing the permeability issues in whole-cell biocatalysis. Significant rate acceleration could be achieved by simple genetic modifications. An order of magnitude increase in reaction was often observed in mutants. The method is general as it has been shown effective with a range of enzyme-substrate systems. Problems with product permeability limitation can be addressed equally well with the approach. The further generalization of the approach by lpp deletion makes this method far more attractive than other permeabilization methods as it can easily be implemented in any E. coli strain of interest, regardless its genetic background. Additionally, the new permeabilization method is more effective than common chemical permeabilization, it achieves its effectiveness with little adverse effects on cell physiology. Thus, it is far more superior to common chemical methods. Furthermore, the method may be applicable for other gram-negative bacteria as lpp homologs are known to exist.
References [1]
[2]
Duetz WA, Beilen JB van, Witholt B. (2001). Using proteins in their natural environment: potential and limitations of microbial whole-cell hydrosylations in applied biocatalysis. Curr Opin Biotech, 12: 419-425. Faber K. (1995). Biotransformations in Organic Chemistry-A textbook, 2nd edition. New York: Springer-Verlag.
Accelerating Whole-cell Biocatalysis by Cellular Membrane Engineering
[3]
[4]
[5]
[6]
[7]
[8] [9] [10] [11] [12]
[13] [14]
[15]
[16] [17]
[18]
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Fontanille P, Larroche C. (2003). Optimization of isonovalal production from α-pinene oxide using permeabilized cells of Pseudomonas rhodesiae CIP 107491. Appl Microb Biotechnol, 60: 534-540. Doig SD, Simpson H, Alphand V, Furstoss R, Woodley JM. (2003). Characterization of a recombinant Escherichia coli TOP10 [pQR239] whole-cell biocatalyst for stereoselective Baeyer–Villiger oxidations. Enzyme Microb Technol, 32:347-355. Michielsen MJF, Meijer EA, Wijffels RH, Tramper J, Beeftink HH. (1998). Kinetics of D-Malate Production by Permeabilized Pseudomonas Pseudoalcaligenes. Enzyme Microb Technol, 22:621-628. De Smet MJ, Wynberg H, Witholt B. (1981). Synthesis of 1,2-epoxyoctane by Pseudomonas olevorans during growth in a two-phase system containing high concentrations of 1-octene. Appl Envrion Microb, 42:811-816. Kondo A, Liu Y, Furuta M, Fujita Y, Matsumoto T, Fukuda H. (2000). Preparation of high acivity whole cell biocatalyst by permeabilization of recombinant flocculent yeast with alcohol. Enzyme Microbial Technol, 27: 806-811. Dupont C, Clarke AJ. (1991). In-vitro synthesis and O acetylation of peptidoglycan by permeabilized cells of Proteus mirabilis. J Bacteriol, 3: 4618-4624. Neidhardt FC, Ingraham JL, Schaechter M. (1990). Physiology of the Bacterial Cell. Sunderland, Massachusetts: Sinauer Associates. Chen. RR. (2007). Permeability issues in whole-cell bioprocesses and cellular membrane engineering. Appl Microbiol Biotechnol, 74: 730-738. Prescott LM, Harley JP, Klein DA. (2002). Microbiology, 5[th] edition. New York: McGraw-Hill Companies, Inc. Vuorio V, Vaara M. (1992). The lipid A biosynthesis mutation lpxA2 of Escherichia coli results in drastic antibiotic supersusceptibility. Antimicrob Agents Chemother, 36: 826-829. Vaara M. (1993). Antibiotic-supersusceptible mutants of Escherichia coli and Salmonella typhimurium. Antimicrobial agents Chemother, 37:2255-2260. Vaara M, Nurminen M. 1999. Outer membrane permeability barrier in Escherichia coli mutants that are defective in the late acyltransferases of lipid A biosynthesis. Antimicrob Agents Chemother, 43: 1459-1462. Plesiat P, Vaara M. (1999). Outer membrane permeability of the antibioticsupersusceptible lipid A mutants of Escherichia coli to hydrophobic steroid probes. J Antimicrob Chemother, 43: 608-610. Yem DW, Wu HC. (1978). Physiological characterization of an Escherichia coli mutant altered in the structure of murein lipoprotein. J. Bacteriol, 133:1419-1426. Lehrer RI, Barton A, Ganz T. (1988). Concurrent assessment of inner and outer membrane permeabilization and bacteriolysis in E. coli by multiple-wavelength spectrophotometry. J Immunol Methods, 108:153-158. Sroga GE, Dordick JS. (2002). Strategy for in vivo screening of subtilisin E reaction specificity in E. coli periplasm. Biotechnol Bioeng, 78:761-769.
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[19] Zylstra GJ, Gibson DT. (1989). Toluene degradation by Pseudomonas putida F1. Nucleotide sequence of the todC1C2BADE genes and their expression in Escherichia coli. J Biol Chem, 264: 14940-14946. [20] Ni Y, Chen RR. (2004). Accelerating whole-cell biocatalysis by reducing outer membrane permeability barrier. Biotechnol Bioeng, 87: 804-811. [21 ]McGregor DR, Elliker PR. (1958). A comparison of some properties of strains of Pseudomonas aeruginosa sensitive and resistant to quaternary ammonium compounds. Can J Microbiol, 4: 499-503. [22] Ni Y, Chen RR. (2005). Lipoprotein mutation accelerates substrate permeabilitylimited toluene dioxygenase-catalyzed reaction. Biotechnol Prog, 21:799-805. [23] Ni Y, Mao Z, Chen RR. (2006). Outer membrane mutation effects on UDP–glucose permeability and whole-cell catalysis rate. Appl Microbiol Biotechnol, 73:384-393. [24] Chen X, Zhang W, Wang J, Fang J, Wang PG. (2000). Production of α-galactosyl epitopes via combined use of two recombinant whole cells harboring UDP-galactose 4epimerase and α-1,3-galactosyltransferase. Biotechnol Prog 16: 595 – 599. [25] Ni Y, Reye J, Chen RR. (2007). Lpp deletion as a permeabilization method. Biotechnol Bioeng, 97:1347-1356. [26] Dumas DP, Caldwell SR, Wild JR, Raushel FM. (1989). Purification and properties of the phosphotriesterase from Pseudomonas diminutia. J Biol Chem, 264: 19659-19665. [27] Donarski WJ, Dumas DP, Heitmeyer DP, Lewis VE, Raushel FM. (1989). StructureActivity Relationships in the Hydrolysis of Substrates by the Phosphotriesterase from Pseudomonas diminuta. Biochemistry, 28: 4650-4655. [28] Shimazu M, Mulchandani A, Chen W. (2001). Cell Surface Display of Organophosphorus Hydrolase Using Ice Nucleation Protein. Biotechnol Prog, 17: 7680. [29] Kang DG, Lim GB, Cha HJ. (2005). Functional periplasmic secretion of organophosphorous hydrolase using the twin-arginine translocation pathway in Escherichia coli. J Bacteriol, 118: 379–385. [30] Castellar MR, Cánovas M, Kleber HP, Iborra JL. (1998). Biotransformation of D(+)carnitine into L(-)-carnitine by resting cells of Escherichia coli O44 K74. J Appl Microbiol, 85: 883-890. [31] Cánovas M, Torroglosa T, Iborra JL. (2005). Permeabilization of Escherichia coli cells in the biotransformation of trimethylammonium compounds into L-carnitine. Enzyme Microb Technol, 37:300-308. [32] Eichler K, Schunck WH, Kleber HP, Mandrand-Berthelot MA. (1994). Cloning, nucleotide sequence, and expression of the Escherichia coli gene encoding carnitine dehydratase. J Bacteriol, 176:2970-2975. [33] Kleber HP. (1997). Bacterial carnitine metabolism. FEMS Microbiol Lett, 147:1-9. [34] Obón JM, Maiquez JR, Cánovas M, Kleber HP, Iborra JL. (1999). High-density Escherichia coli cultures for continuous L-carnitine production. Appl Microbiol Biotechnol, 51: 760-764.
In: Biocatalysis Research Progress Editors: F. H. Romano, A. Russo
ISBN: 978-1-60456-619-2 © 2008 Nova Science Publishers, Inc.
Chapter X
Recent Advances in Enzymatic Synthesis of Water-Soluble Conducting Polymers Estibalitz Ochoteco*, Tomasz Sikora, David Mecerreyes, Jose A. Pomposo, and Hans Grande New Materials Department, CIDETEC-Centre for Electrochemical Technologies, Parque Tecnológico de San Sebastián, Paseo Miramón 196, E-20009 Donostia-San Sebastian, Spain, Email:
[email protected] Phone: 34943309022; Fax: 34943309136
Abstract The present chapter introduces the newest synthesis strategies developed by our research group in order to overcome the main disadvantages derived from enzymatic biocatalysis as a strategy for producing conducting polymers. First, the enzymatically catalyzed polymerization of 3,4-ethylenedioxythiophene (EDOT) in the presence of polystyrenesulfonate (PSS) is achieved showing an electrical conductivity of 2×10-3 S/cm and an excellent film formation ability as confirmed by atomic force microscopy images. Second, a simple method for immobilizing horseradish peroxidases (HRP) in the biocatalytic synthesis of polyaniline (PANI) is presented. This method is based on a biphasic catalytic system where the enzyme is encapsulated inside the ionic liquide (IL) 1-butyl-3-methylimidazolium hexafluorophosphate, while other components remain in the aqueous phase. The enzyme is easily recovered after reaction and reused several times. Finally, a new bifunctional template (sodium dodecyl diphenyloxide disulphonate, DODD) is proposed in the synthesis of polyaniline (PANI) as a strategy to improve water solubility as well as electrical conductivity in the obtained polymer.
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Introduction The field of intrinsically conducting polymers has attracted great attention due to their interesting electrical and optical properties. Since their discovery in 1977, they have been investigated for countless technological applications, such as organic lightweight batteries, microelectronics, optical displays, antistatic coatings, and electromagnetic shielding [1]. These extremely promising materials have traditionally been synthesized by monomer oxidation in the presence of a strong oxidant showing high insolubility and intractability when synthesized [2]. In this sense, the use of enzymes as biocatalysts for the synthesis of conducting polymers has been studied in the last years as a “green synthesis process” alternative [3]. Enzymes can offer environmentally benign reaction conditions [4], a high yield of polymerization, and superior level of control in regioregularity and stereochemistry [5], consequently resulting in soluble and processable conducting polymers [6]. In biocatalytic reactions, a peroxidase enzyme such as horseradish peroxidase (HRP) or others such as palm tree peroxidase or laccasse are used as catalyst for the aniline polymerization in an aqueous buffer. As polymer begins to be synthesized, it precipitates out showing processability problems 7]. Although several strategies have been used to avoid this precipitation as modification of monomer structures [8, 9], synthesis in micellar media [10], reverse micelles [11, 12] and interfacial polymerizations [13], the final conductivity of the obtained PANI is poor in most cases. Liu and coworkers [14, 15] improved successfully the conductivity of enzymatic synthesized PANI by using a polymer electrolyte such as polystyrenesulphonate (PSS) in the reaction media. This water soluble polyelectrolyte serves as a linear template. Since PSS and aniline have a pKa of 0.70 and 4.63, respectively, at pH close to 4, PSS is anionically charged, and aniline cationically charged. Then, aniline and PSS complex electrostatically so aniline monomers become more ordered and para-directed reaction is favoured, giving a more linear, highly conjugated structure. Consequently, the PANI conductivity is improved by this method. The same effect is reached with dodecylbenzene sulfonic acid (DBSA), hexadecyltrimethylammonium bromide (HDTMAB) and polyoxyethylene isooctyphenyl ether (PEOPE) as templates. These molecules aggregates to form micelles above a critical concentration (critical micellar concentration, CMC), creating another template configuration for a guided polymerization, while the use of small molecules such as sodium benzene sulfonic acid (SBS), without the ability to form these templates does not form a highly conducting PANI by enzymatic synthesis. So, it seems clear that the aggregation of anionic species (via aggregation of small molecules or polymeric electrolytes) is essential to create an adequate environment giving necessary counter-anions for the doping of the conducting polymer, a way to keep water solubility, and an alignment for monomer molecules during polymerization. In spite of these last advances, the electrical conductivity in the final complex remains low for practical applications, and new strategies are still being searched to attain high conductivity and excellent water solubility. One of the most important drawbacks of this synthesis strategy is the cost of the enzyme. In general, for practical applications the expensive enzymes must be recovered and reused after the reaction. This is the main reason behind the well established strategy of immobilization of enzymes into solid supports which was applied to HRP enzyme [16].
Recent Advances in Enzymatic Synthesis of Water-Soluble Conducting Polymers 247 Actually, new alternative strategies are being studied in many research groups for recycling and reusing the enzyme in the biocatalysis synthesis. Finally, it is worth mentioning that the biocatalytic synthesis process has been successfully applied to PANI as conducting polymer, but it has not yet been extended to other technologically interesting polymers such as polythiophenes [17]. Of particular interest, poly(3,4-ethylenedioxithiophene), PEDOT is becoming the most successful for commercial applications. This lack of polymerization success is clearly stated in a very recent paper [18], where this lack is said to be probably due to the high oxidation potential of monomers such as 3,4-ethylenedioxythiophene (EDOT) and pyrrole (Py), referred to the oxidation potential of the HRP/H2O2 pair, concluding that these monomers are inappropriate for this enzymatic synthesis process. In order to provide an answer to the problematic issues detected in the field of biocatalytic synthesis, new synthetic strategies have been developed in our research group very recently. A) Contrary to the statements previously made by other authors, the first enzymatic polymerization of EDOT was achieved, demonstrating that this green synthesis process can be applied to other technologically interesting polymers apart from PANI. B) An ionic liquid based biphasic catalytic system was employed as a mean to recover and to reuse the enzyme several times in aniline or EDOT polymerization. And as a third development, C) new bifunctional templates were used in the enzymatic polymerization giving as a consequence highly soluble and conducting polymers. This chapter introduces and discusses these recent synthetic strategies in the enzymatic biocatalytic approach.
Experimental Section Materials
Horseradish Peroxidase (HRP, EC 1.11.1.7, type II, 150-200 units/mg solid) aniline (99%), and EDOT (99%) were purchased from Sigma-Aldrich Chemical S.A. Sodium dodecyl diphenyloxide disulphonate (DODD) was obtained from Dow Chemicals Co. (Scheme 1b). Hydrogen peroxide (30 wt %) and hydrochloric acid (1 N) were obtained from Quimibacter S.L. The hydrogen peroxide was diluted to 0.3% in deionized water and hydrochloric acid served to adjust the pH of bulk solution before the addition of hydrogen peroxide. The concentration of HRP stock solution was 3 mg/mL.
Enzymatic Polymerization of Aniline in the Presence of DODD
In a typical polymerization reaction, equimolar quantities (typically 0.055 M) of the surfactant template DODD and aniline were dissolved into 20 mL of distilled water at room temperature. Next, the pH was adjusted to 4.1 with addition of HCl. Micelles were formed spontaneously when the concentration of surfactant in the solution was greater than the CMC (critical micellar concentration); the known CMC of DODD is 7 mM [19]. To this solution, 2 mL of HRP solution (3 mg in 1 mL of distilled water) was added under magnetic stirring.
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Finally, an equimolar amount (typically 0.055 M) of diluted hydrogen peroxide solution was added over a period of 12 minutes. The colour of the reaction solution changes with H2O2 addition. The colour sequence is light blue, blue green, green, and more and more dark with H2O2 amount increasing. After the addition of H2O2, the reaction was left under stirring for 3 hours to ensure a complete polymerization process.
Enzymatic Polymerization of EDOT in the Presence of PSS
The synthesis procedure was as the one previously described for the aniline unless in this case, the pH was adjusted to 2, polymerization temperature was 4ºC, and polymerization time 16 hours.
Enzymatic Polymerization of Aniline in a Biphasic Biocatalytic System
At first, we dissolved the HRP enzyme into the IL (6 mg into 1 mL). This IL was added to 10 mL of aqueous solution of aniline, the template dodecylbenzenesulfonic acid (DBSA) and hydrogen peroxidase at pH 4.3. In the basic polymerization, equimolar quantities (typically 0.055 M) of the template, aniline and hydrogen peroxide were dissolved. To use DBSA like a template is necessary that concentration is greater than its CMC (Critical Micellar Concentration) around of 1.6*10-3 M. Reaction was carried out at 20 ºC for 3 hours. After reaction, the IL phase was separated from the aqueous phase by liquid/liquid pour off in order to purify the PANI aqueous solution and to recuperate the HRP/IL phase.
Characterization Methods
UV-vis spectra were recorded on a UV-1603 SHIMADZU spectrometer. In each measurement distilled water was used as control. The electrical conductivity of PANI/DODD, PANI/DBSA, and PEDOT/PSS samples was measured with a home-made four-probe instrument. Micelle particle sizes were measured using a Beckman Coulter N5 Submicron Particle Size Analyzer. The surface morphology was obtained by a Pico Plus Atomic Force Microscope operating in acoustic mode.
Results and Discussion First enzymatic Polymerization of EDOT
The biocatalytic synthesis process has been successfully applied to polyaniline as conducting polymer, but it has not yet been applied to other technologically interesting polymers such as polythiophenes17. In a previous work by other researchers18 this lack was
Recent Advances in Enzymatic Synthesis of Water-Soluble Conducting Polymers 249 said to be probably due to the high oxidation potential of monomers such as 3,4ethylenedioxythiophene (EDOT) and pyrrole (Py), compared to the oxidant HRP/H2O2, concluding that these monomers are inappropriate for this enzymatic synthesis process. In our laboratory a special attention was paid to the enzymatic polymerization of the EDOT monomer. Surprisingly, our first trials succeeded and a blue colored polymer solution was obtained after 16 hours of reaction (Scheme 1). It is well known that an acidic media is suitable in order to increase the rate of polymerization. The protonic acids and a variety of Lewis acids catalyse the equilibrium reaction of EDOT to the corresponding dimeric and trimeric compounds without further oxidation or reaction [20]. In our case, three different pH were studied (pH=2, pH=4 and pH=6) in order to establish the most optimum one for the adequate synthesis of EDOT. Figure 1 shows the UV visible spectra for these three reactions. As can be observed, the only spectra showing the presence of bipolaron absorption band at 800 nm associated to PEDOT polymer was the one obtained at pH = 2.
1
Absorbance
0,8
0,6
0,4
0,2
0 200
400
600
800
1000
1200
Wavelength (nm)
Reprinted with permission from Rumbau V. et al., Biomacromolecules, 2007, 8(2), 315 [21]. © 2007 American Chemical Society. Figure 1. UV-vis spectra for PEDOT complexes synthesized at different pH for 16 hours. a) pH=2, b) pH=4, pH=6.
Our results demonstrate that the high oxidation potential of the monomer may not be the reason of the lack of polymerization in previous failed attempts. Water soluble PEDOT can be easily obtained if appropriate working conditions are used (pH=2, and a polymerization time as long as long as 16 hours). On the other hand, as it is well known, the activity of the HRP enzyme decreases abruptly in this acidic media (in pH=4, the HRP activity is around 0 after 1 hour1b) but, surprisingly, even if the enzymatic activity should normally be finished sometime after the first 60 minutes, the reaction was observed to proceed uninterruptedly
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throughout the 16 hours. These unexpected results led us to think that under these strong acidic conditions the enzyme could be deactivated and the polymerization process could be triggered only by H2O2. Thus, a control experiment was performed employing the same described conditions, with no HRP enzyme. Since, in this case, no polymerization was observed, the HRP enzyme was assumed to be the catalyst triggering the polymerization reaction. The explanation for the enhanced enzyme activity proposed by the authors is the presence of an excess of EDOT monomer in the media. As the solubility of the monomer is very poor in water, the reaction takes place in a biphasic media, where the HRP is localized more preferentially in the monomer phase. To prove this statement, a mixture of EDOT, template and enzyme was prepared in acidic water (pH=2). This solution was kept under magnetic stirring for 24 hours. Then, a solution of H2O2 was added. The reaction proceeded as usual and a conducting water soluble PEDOT was obtained after additional 16 hours. Thus, the EDOT monomer droplets would act as a) enzyme protectors against deactivation, as well as b) monomer feed to keep a constant EDOT concentration in the reaction media. As conclusion, the enzymatically catalyzed polymerization of EDOT in presence of PSS can successfully be performed, if specific synthesis conditions are employed (low pH, protected enzyme). The obtained PEDOT polymer shows a very good film formation capacity at a macroscopic level, and very low surface roughness also at a microscopic level, as can be observed in the surface picture of the film obtained by atomic force microscopy (Figure 2).
Reprinted with permission from Rumbau V. et al., Biomacromolecules, 2007, 8(2), 315 [21]. © 2007, American Chemical Society. Figure 2. Atomic Force Microscopy pictures obtained from a PEDOT/PSS film (6μ×6μ) synthesized at a EDOT/PSS ratio of 1:1.
Recent Advances in Enzymatic Synthesis of Water-Soluble Conducting Polymers 251 A Biphasic Biocatalytic System for the Synthesis of Conducting Polymers: Enzyme Recovery
In general, for practical applications the expensive enzymes must be recovered and reused after the reaction. This is the main reason behind the well established strategy of immobilization of enzymes into solid supports which was applied to HRP enzyme16. As an alternative strategy, our laboratory reported a new method allowing to recycle and reuse the HRP enzyme in the biocatalytic synthesis, in this case, of PANI, but it could be easily extended to other Intrinsically Conducting Polymers (ICPs). The method was based on the use of a biphasic catalytic system where the enzyme was encapsulated by simple solubilisation into an ionic liquid (IL). The main strategy consisted of encapsulating the HRP enzyme in room-temperature ILs insoluble with water, and the other components of the reaction such as aniline monomer, H2O2 oxidant and DBSA template in the aqueous phase. The biocatalytic process should take place at the aqueous/IL interface, and the HRP enzyme could be subsequently recovered by liquid/liquid phase separation after the biocatalytic reaction (Figure 3).
Stirring
Reaction
Phase separation
aniline H2O2 DBSA
Recovered IL Phase+enzyme Aqueous Phase/ PANI IL Phase+enzyme Reprinted with permission from Rumbau V. et al., Macromolecules, 2006, 39(25), 8547 [22]. © 2006, American Chemical Society. Figure 3. Biphasic biocatalytic system in conducting polymers synthesis.
After 3 hours of reaction, the UV-vis spectra of the obtained aqueous phase was compared to the PANI obtained by classical biocatalytic synthesis dissolving the HRP enzyme in the aqueous reaction media (Figure 4). Both spectra were similar and clearly showed an absorption peak maximum at 420 nm. Furthermore, a well known absorption tail extended towards the near-IR region between 800-1100 nm arising from the polaron population in the material which is a sign of the formation of conducting PANI.
Estibalitz Ochoteco, Tomasz Sikora, David Mecerreyes, et al.
Absorbance a.u.
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Reprinted with permission from Rumbau V. et al., Macromolecules, 2006, 39(25), 8547 [22]. © 2006, American Chemical Society. Figure 4. UV-Vis spectra of PANI/DBSA aqueous solution obtained without encapsulation of the HRP enzyme (upper spectrum) and with encapsulation of the HRP enzyme in 1-butyl-3-methylimidazolium hexafluorophosphate (lower spectrum).
The recovered HRP/IL mixture was added to a new aqueous reaction media, including fresh aniline, DBSA and H2O2. A second reaction took place and the HRP/IL was recovered again by liquid-liquid phase separation. The same process was successfully repeated up to 5 times using the same HRP/IL phase. The electrical conductivity of the PANI films prepared by solvent casting from the aqueous solutions showed a relatively high and similar value even after the fifth run (Figure 5) demonstrating the validity of our approach and the ease recyclability and reuse of the enzyme inside the IL. -1
Log s (S/cm)
-2 -3 -4 -5 -6 -7 0
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Reprinted with permission from Rumbau V. et al., Macromolecules, 2006, 39(25), 8547 [22]. © 2006 American Chemical Society. Figure 5. Electrical conductivity films obtained from the aqueous solutions after several runs. Run 0 indicates the control value obtained without encapsulation of the enzyme.
Recent Advances in Enzymatic Synthesis of Water-Soluble Conducting Polymers 253 New Bifunctional Templates as a Mean to Improve Electrical Conductivity
As previously stated, the use of templates as DBSA, HDTMAB and PEOPE allows the formation of well-defined micellar structures in aqueous solution when the concentration is over its critical micellar concentration (CMC). In this case, our group supposed that polymerization with DODD as template proceeded by a micellar mechanism in the same way. In an aqueous acid solution of DODD and aniline, anilium ions locate at the micellar interface with benzene parts penetrating into the hydrophobic core of the DODD micelle to form the complex as illustrated in Scheme 2b. At the concentration of 0.055 mol/l a slight turbidity was observed in solution indicating a micellar aggregation. Once the micellar structure is formed and the enzyme is added to the aqueous media, addition of H2O2 triggers the polymerization of anilinium ions around micelles (Figure 6). The bifunctional DODD molecule, due to the presence of two sulphonic groups in its structure, shows a higher water solubility than monofunctional surfactants such as DBSA. At acidic pHs sulphonate groups become negatively charged increasing total charge per molecule, and consequently, solubility. This idea is reflected in the fact that CMC value for DODD is higher than DBSA’s one (7×10-3 and 1.6×10-3 M, respectively). Furthermore, in a bifunctional surfactant each micelle is expected to be composed by a lower number of individual molecules than the ones involved in a micelle created by monofunctional ones, and consequently, to be smaller in size. Light scattering measurement verified this assumption, where micelles of Ø < 4 nm and Ø=5 nm in size were observed for DODD and DBSA, respectively. We can assume, therefore, that for a same surfactant concentration, there are more micelles for a bifunctional surfactant than for a monofunctional one. The content of anionic charges, and the anionic template surface for complex formation in the media should, consequently, be higher. It is well known that the solubility of the polymer-complex is explained by the anionic charges on the template, and that a percentage of these anionic charges must remain free to keep the complex in solution. Our system being composed by a higher anionic surface, the complexation with anilinium cations on the surface will be higher, and consequently higher the polymerization conversion before precipitation occurs. This higher local anilinium cation concentration should give to higher molecular weights and, consequently, higher conductivity in the product. Experimental results (Figure 7) show that for PANI/DODD and PANI/DBSA synthesized at the same conditions, the conductivity of he PANI/DODD complex is higher than that of the PANI/DBSA. Furthermore, the DODD template allows the use of a higher aniline/template ratio before precipitation occurs. PANI/DBSA complexes show precipitaction at ratios close to 2.5/1 whereas PANI/DODD complexes remain in solution even at a ratio of 4/1.
NH2
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[DODD]>CMC
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O S O
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O O +
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S O
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S O
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NH3
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NH3
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NH3
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NH3
S O
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+
NH3
O
S
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O
O
S
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O
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S O
+
O
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O O O S
O
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S
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HRP H2O2
O +
Coupling
NH3 O O
O
S O
O
O
S
O H3N
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O S
O
O S
O
O
+
O
H
O
*
O S O O
O O
O
O H3N
S
+
O
O
O
O
+
O
NH3
-O3S
+
N
-O3S
NH3
S O
O S O
N
S O O
H
+
N H
S O
O
Aniline/DODD micelles interactions
Reprinted with permission from Rumbau V. et al., Enzyme and Microbial Technology, 2007, 40(5), 1412 [23]. © 2007, Elsevier. Figure 6. Polymerization mechanism: aniline-DODD interaction and polymerization.
N n
+
O
H
PANI/DODD
*
Recent Advances in Enzymatic Synthesis of Water-Soluble Conducting Polymers 255
Conductivity (S/cm )
0,06
0,04
PANI/DODD 0,02
PANI/DBSA
0 0
1
2
3
Ratio (Aniline/Template) Reprinted with permission from Rumbau V. et al., Enzyme and Microbial Technology, 2007, 40(5), 141223. © 2007, Elsevier. Figure 7. Influence of aniline/template ratio on conductivity for (♦) enzymatic PANI/DODD and (■) enzymatic PANI/DBSA.
Conclusion New strategies were successfully introduced in the field of biocatalytic synthesis of ICPs in order to overcome some of the problematic issues of this technology for practical applications. The enzimatically catalyzed polymerization of EDOT in presence of PSS was successfully performed, proving that the biocatalytic strategy can be also applied to technologically interesting conducting polymers others than PANI. The obtained PEDOT polymer films showed excellent electrical conductivity (2×10-3 S/cm), as well as film formation capacity, with excellent surface roughness at microscopic level. A new biphasic polymerization strategy was demonstrated allowing the purification of the obtained conducting polymer aqueous solution from the remaining enzyme, while the recovered enzyme can be recycled and reused several times. Finally, a new bifunctional template was selected and used in the enzymatic synthesis of PANI (DODD), reaching, as a consequence, higher solubilities and electrical conductivities than the ones obtained when monofunctional templates were used.
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References Nagarajan, R.; Tripathy, S.; Kumar, J.; Bruno, F.F.; and Samuelson, L. An Enyzmatically Synthesized Conducting Molecular Complex of Polyaniline and Poly(vinyl phosphonic acid). Macromolecules, 2000, 33, 9542. b) Liu, W., Kumar, J., Tripathy, S., Senecal, K.J., and Samuelson, L. Enzymatically synthesized conducting polyaniline", W. Liu, J. Kumar, S. Tripathy, K.J. Senecal and L. Samuelson. J. Am. Chem. Soc. 1999, 121(1), 71. [2] Annis, B.K.; Narten, A.H.; MacDiarmid A.G.; and Richter, A.F. A. covalent bond to bromide in HBr-treated polyaniline from. X-ray diffraction. Synt. Met. 1988, 22, 191. [3] Thiyagarajan, M.; Samuelson, L.A.; Kumar, J.; and Cholli, A.L. Helical Conformational Specificity of Enzymatically Synthesized Water-Soluble Conducting Polyaniline Nanocomposites. J. Am. Chem. Soc. 2003, 125, 11502. b) Wang, X.; Schreuder-Gibson, H.; Downey, M.; Tripathy, S.; and Samuelson, L. Conductive fibers from enzymatically synthesized polyaniline. Synthetic Metals. 1999, 107, 117. c) Liu, W.; Cholli, A.L.; Nagarajan, R.; Kumar, J.; Tripathy, S.K.; Bruno, F.F.; and Samuelson L. Enzymatically Synthesized Conducting Polyaniline. J. Am. Chem. Soc. 1999, 121, 11345. d) Liu, W.; Cholli, A.L.; Kumar, J., Tripathy, S.; and Samuelson, L. Mechanistic Study of the Peroxidase-Catalyzed Polymerization of Sulfonated Phenol. Macromolecules, 2001, 34, 3522. [4] [4] Cruz-Silva, R.; Romero-García, J.; Angulo-Sanchez, J.; Ledezma-Perez, J.; AriasMartín, E.; Moggio, I.; and Flores-Loyola, E. Template-free enzymatic synthesis of electrically conducting polyaniline using soybean peroxidase. Eur. Pol. J. 2005, 41, 1129. [5] Lim, C.H. and Yoo, Y.J. Proc. Biochem. 2000, 36, 233. [6] Xu, P.; Singh, A. and Kaplan, D.L. Enzymatic catalysis in the synthesis of polyanilines and derivatives of polyanilines. Enzyme Catal Synth Polym. 2006, 194, 469. [7] Saunders, B.C.; Holmes-Siedle, A.G.; and Stark, B.P. The properties and uses of a versatile enzyme and of some related catalysts. In: Peroxidase. London: Butterworths; 1964, 214. [8] Alva, K.S.; Kumar, J.; Marx, K.A.; and Tripathy, S.K. Enzymatic synthesis and characterization of a novel water-soluble polyaniline: poly(2,5diaminobenzenesulfonate). Macromolecules, 1997, 30, 4024. [9] Alva, K.S.; Marx, K.A.; Kumar, J.; and Tripathy, S.K. Biochemical synthesis of water soluble polyanilines: poly(p-aminobenzoic acid). Macromol Rapid Commun. 1996, 17, 859. [10] Liu, W.; Wang, J.D.; Ma, L.; Liu, X.H.; Sun, X.D.; Cheng, Y.H. et al. Enzymatic polymerization of p-phenylphenol in aqueous micelles. Ann NY Acad Sci. 1995, 750, 138. [11] Rao, A.M.; John, V.T.; Gonzalez, R.D.; Akkara, J.A.; and Kaplan, D.L. Catalytic and interfacial aspects of enzymatic polymer synthesis in reversed micellar systems. Biotechnol Bioeng. 1993, 41, 531. [1]
Recent Advances in Enzymatic Synthesis of Water-Soluble Conducting Polymers 257 [12] Premachandran, R.S.; Banerjee, S.; John, V.T.; McPherson, G.L.; Akkara, J.A.; and Kaplan, D.L. The enzymatic synthesis of thiol-containing polymers to prepare polymerCdS nanocomposites. Chem Mater. 1997, 9, 1342. [13] Bruno, R.; Akkara, J.A.; Samuelson, L.A.; Kaplan, D.L.; Marx, K.A.; Kumar, J. et al. Enzymatic mediated synthesis of conjugated polymers at the Langmuir trough air– water interface. Langmuir. 1995, 11, 889. [14] Samuelson, L.A.; Anagnostopoulos, A.; Alva, K.S.; Kumar, J.; and Tripathy, S.K. Biologically Derived Conducting and Water Soluble Polyaniline. Macromolecules. 1998, 31, 4376. [15] Liu, W.; Kumar, J.; Tripathy, S.; Senecal, K.J.; and Samuelson, L.A. Enzymatically Synthesized Conducting Polyaniline. Am. Chem. Soc. 1999, 121(1), 71. [16] Jim, Z.; Su, Y. and Duan, Y. A novel method for polyaniline synthesis with the immobilized horseradish peroxidase enzyme. Synth. Met. 2001, 122, 237; b) Fernandes, K.F.; Lima, C.S.; Pinho, H.; and Collins, C.H.; Proc. Biochem. 2003, 38, 1739. c) Fernandes, K.F.; Lima, C.S.; Lopes, F.M.; and Collins, C.H. Properties of horseradish peroxidase immobilised onto polyaniline Proc. Biochem. 2004, 39, 957. d) Azevedo, A.M.; Vojinovic, V.; Cabral, J.M.S; and Gibson, T.D.; Fonseca, L.P. Operational stability of immobilised horseradish peroxidase in mini-packed bed bioreactors. J. Mol. Cat. B: Enzym. 2004, 28, 121. e) Sun, D.; Cai, C.; Li, X.; Xing, W.; and Lu, T. Direct electrochemistry and bioelectrocatalysis of horseradish peroxidase immobilized on active carbon. J. Electroanal, Chem. 2004, 566, 415. f) Moeder, M.; Martin, C. and Koeller, G. Degradation of hydroxylated compounds using laccase and horseradish peroxidase immobilized on microporous polypropylene hollow fiber membranes. J. Memb. Sci. 2004, 245, 183. [17] Bruno, F.F.; Akkara, J.A.; Kaplan, D.L.; Sekher, P.; Marx, K.A.; and Tripathy, S.K. Novel enzyme-mediated two-. dimensional polymerization of aromatic derivatives on. a Langmuir Trough. Ind. Eng.Chem. Res. 1995, 34, 4009. (b) MacDiarmid, A.G. and Epstein, A.J. Application of Thin Films of Polyaniline and Polypyrrole in Novel LightEmitting Devices. ACS Symp. Ser. 1997, 672, 395. (c) Skotheim, T.A.; Elsenbaumer, R.L.; and Reynolds, J.R. Handbook of Conducting Polymers, Marcel Dekker: New York, 1998. d) Bruno, F.F; Nagarajan, R.; Roy, S.; Kumar, J.; Samuelson, L.A. Biomimetic Synthesis of Water Soluble Conductive Polypyrrole and Poly(3,4Ethylenedioxythiophene). Journal of Macromolecular Science: Part A-Pure and Applied Chemistry. 2003, A40(12), 1327. [18] Bruno, F.F.; Fossey, S.A.; Nagarajan, S.; Nagarajan, R.; Kumar, J.; and Samuelson, L.A. Biomimetic synthesis of water-soluble conducting copolymers/homopolymers of pyrrole and 3,4-ethylenedioxythiophene. Biomacromolecules, 2006, 7, 586. [19] Aramendia, E.; Barandiaran, M.J.; Grade, J.; Blease, T.; and Asua, J.M. Polymerization of high-solids-content acrylic latexes using a nonionic polymerizable surfactant. Journal of Polymer Science: Part A: Polymer Chemistry. 2002, 40, 1552. [20] Kirchmeyer, S. and Reuter, K. Scientific importance, properties and growing applications of poly(3,4-ethylenedioxythiophene). J. Mater. Chem. 2005, 15, 2077.
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[21] Rumbau, V.; Pomposo, J.A.; Eleta, A.; Rodriguez, J.; Grande, H.; Mecerreyes, D.; and Ochoteco, E. First Enzymatic Synthesis of Water-Soluble Conducting Poly(3,4ethylenedioxythiophene). Biomacromolecules, 2007, 8(2), 315. [22] Rumbau, V.; Marcilla, R.; Ochoteco, E.; Pomposo, J.A.; and Mecerreyes, D. Ionic Liquid Immobilized Enzyme for Biocatalytic Synthesis of Conducting Polyaniline. Macromolecules, 2006, 39(25), 8547. [23] Rumbau, V.; Pomposo, J.A.; Alduncín, J.A.; Grande, H.; Mecerreyes, D.; and Ochoteco, E. A new Bifunctional Template for the Enzymatic Synthesis of Conducting Polyaniline. Enzyme and Microbial Technology, 2007, 40(5), 1412.
In: Biocatalysis Research Progress Editors: F. H. Romano, A. Russo
ISBN: 978-1-60456-619-2 © 2008 Nova Science Publishers, Inc.
Chapter XI
Marine Enzymes for Biocatalysis – Production, Isolation and Applications K. Muffler1, J. Mukherjee2 and R. Ulber1 1
University of Kaiserslautern, Department of Mechanical and Process Engineering, Gottlieb-Daimler-Straße, 67663 Kaiserslautern 2 Environmental Science Programme and Department of Life Science and Biotechnology, Jadavpur University, Kolkata 700 032, India
Abstract The diversity of marine life offers a variety of novel enzymes which might have a tremendous potential as biocatalysts for academic research as well as for industrial processes. Concerning the wide spectrum of ecological habitats, which differs extremely in temperature, pressure, and salinity, the whole machinery of enzyme expression and stability of the proteins was adopted to the distinct environmental conditions of the producing micro- as well as macroorganisms. Therefore the oceans provide an almost untapped reservoir of biocatalysts showing interesting properties like high salt, pressure, and temperature tolerance. With respect to the special requirements of industrial processes which are particularly focused on high mass transfer and high time space yields, respectively, enzymes from marine origin located at exotically regions can help to fulfill these specific demands. Thus, applications of such proteins or whole cells as catalysts for manufacturing bulk- and fine chemicals seems visionary up to know. However, regarding the current trends and developments in molecular biology as well as genetic engineering it appears more realistic in the case of long-term view. By means of identifying the origin of the enzymes’ stability one has the possibility to modulate other so far unstable (terrestrial) enzymes. This review covers the development and research work done on the processing of enzymes from marine origin, which can be used as biocatalyst tools for research in academia and industry.
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1. Introduction A characteristic trait of the marine habitat is its tremendous size and its vast biodiversity, which is represented in distinct habitats such as deep-sea, coral reefs, tropical or artic and hydrothermal zones, respectively. Although life evolved exclusively in the marine realm within the Precambrian 3.5 billion years ago and considering the fact, that the oceans are currently covering more than 70% of the earth’s surface, biosphere research and screenings for natural products were mainly focused for a long time on terrestrial habitats, especially on the Streptomyces genus [1]. Within the screening of novel pharmaceutical lead structures the research extends its scope also on marine micro- as well as macroorganisms approximately 30 years ago. The success of this new focus is obviously, since more than 10,000 novel compounds were detected within the last two decades [2,3]. However, it is assumed that novel pharmacological relevant compounds are exclusively from marine origin [4]. These screening activities were in its initial phase - mainly driven by the development of novel bioactive compounds rather than novel marine enzyme systems. By now the natural product research focuses also on the enzyme systems necessary for the production of the bioactive secondary metabolites, as for example polyketide synthases [5,6], catalyzing the linkage of acyl-coenzyme A subunits via generation of cyclic structures similar to well known bioactive polyketides as bryostatin 1. Aside from the natural product research some (novel) marine enzymes, catalyzing a multitude of processes relevant for biotransformations in White Biotechnology, showing astonishing properties like high salt tolerance, hyperthermostability, barophilicity and cold adaptivity. Their performance results from the adaptation of their host organisms during the evolutional development to the environmental surrounding. However, the raising demand for environmental friendly and beneficial economical manufacturing of bulk and fine chemicals requires the exploitation of these enzymes for the specific and selective production of high value compounds like chiral amines, alcohols, halogenated amino acids as well as the enzymes itself in the case of for example thermostable proteases or polymerases. Up to now industrial biotransformations are often performed by application of the Corynebacterium glutamicum enzyme system set-up, but marine organisms offer an enormous source of new or more active biocatalysts. The introduction of such enzymes into industrial-scale approaches of biotransformations necessitates a sustainable access of the applied enzymes from marine origin, if one considers the difficulties of enzyme expression or cultivation of the natural expression system, when transferred from its originally habitat to laboratory conditions. This approach includes identification, isolation, characterization and cloning of the responsible genes in adequate host organisms as well as optimization of enzyme stability and enzyme expression. Aside from such microorganisms as bacteria or fungi also higher organisms such as fishes, prawns, crabs, snakes, plants, and algae can be used for tapping of marine enzymes [7]. Especially the latter one accounts for a number of interesting halogenating enzymes species. The next chapters give a deeper insight into handling and cultivation of the marine enzymes’ host concerning the expression of the biocatalysts. Additionally, several industrial significant examples of enzymes from marine origin for biotransformation processes are presented, while in the main focus are starch degrading enzymes, proteolytic enzymes, DNA
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polymerases, and halogenating enzymes. Since the downstream process is presumably the most underestimated step within a bioprocess, the last chapter of this review is dedicated current purification techniques for proteins from cell debris and extracellular expressed proteins from fermentation broths of marine organims. Such a process is exemplified concerning a marine derived sulfite-oxidizing enzyme, which can be used as a beneficial tool for establishing a sulfite biosensor.
2. Sources and Handling of Marine Enzymes Marine enzymes can be anyway isolated from each organism which derives from marine habitats. However, the microorganismal pool certainly represents the most investigated group of the marine ecosystem, especially bacteria, fungi and extremophiles rather than higher macroorganisms. These microorganisms can be found within the free water column, in marine sediments or in symbiotic associations (endosymbiontic and surface associated) with higher marine organisms. Prior to examination of enzyme expression, these microorganisms must be successfully cultivated in the laboratory. However, a transfer of the marine organisms from their natural ecosystems to laboratory conditions often failures. The principal reason is presumably the adaptation of the organism to its usual surrounding and its participation on several symbiotic interactions with other organisms, which is usually not considered by the experimenter. A crucial factor allowing a more or less effective and reproducible cultivation of organisms from marine origin is to provide as far as possible similar environmental conditions to the marine strain in laboratory cultivations. It is obvious that microorganisms deriving from the free water column have distinct requirements concerning the nutrient concentration than such organisms which are growing on surfaces or within the tissue of e.g. invertebrates. While sea water contains approximately 104-106 cells per ml much higher cell densities of six orders of magnitude can be detected on surfaces exposed to sea water [8,9]. Since sea water can be regarded as a nutrient poor medium, it is assumed that free living microorganisms have adapted themselves to these conditions. Commonly applied media for fermentation of marine microorganisms have a high carbon content, which does not reflect the specific nutrient requirements of the strains. While application of the so-called dilution technique several strains by then classified as uncultivatable were assessable for fermentation under laboratory conditions [10]. However, the growth of marine bacteria is not only limited to the carbon content of sea water but also limited with regard to iron availability [11,12]. Since such a limited bioavailability of iron is ubiquitous several organisms have adapted to these conditions and are capable to release special iron-complexing agents, the so-called siderophores, which enables the organism to increase iron-resorption [13]. However, for cultivation of marine microorganisms under lab-scale conditions many experiments were carried out by application of the so-called Marine Broth 2216, which can be regarded as a well established reference medium. To reflect the special nutrient needs of distinct marine microorganisms it is often utilized in diluted concentrations. To satisfy the need of iron, it contains a well accessible iron salt (0.1% iron-citrate).
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Table 1. Mass transfer of several molecules in biofilms, represented by the ratio of diffusion coefficients of biofilm to water, according to [9] Molecule
Molar Mass [g/mol]
Dbiofilm/Dwater
O2
32
0.98
fluorescein
332
0.91
dextran
4.000
0.76
dextran
38.000
0.85
dextran
580.000
0.26
bovine serum albumine
68.000
0.86
catalase
225.000
0.67
DNA
3.200.000
0.63
Since the marine microorganisms utilized for production of marine products such as enzymes as well as natural secondary metabolites are adapted to their naturally surrounding, their exploitation for production purposes is commonly a challenging task. This applies especially, when the organisms derived from sea water exposed surfaces or from tissues of higher marine organisms. Within these environments the microorganisms are commonly grown as conglomerates of bacteria, fungi, algae, and protozoans, respectively. Thus, higher orders of specializations of distinct microorganisms can be observed in such a biofilm. The interactions are regulated via different (chemical) communication patterns. Communication is possible between organisms of the same species as well as different (higher) species, while the chemical compound facilitating the communication varies. Such communication requires a beneficial diffusion of the communication/signal molecule within the matrix. As presented in Table 1 biofilms are featuring beneficial diffusion coefficients for small molecules. While microorganisms are exposed to changing conditions in their natural habitats with regard to pH, temperature, osmolality or nutrients they have evolved specific adaptation systems. Thus, they are capable to activate or repress the expression of target genes in response to environmental conditions [14]. Further mentionable mechanisms which are based on stimuli-response mechanisms can be found in the bacterial cell-cell communication by means of small signal molecules, called autoinducers. Up to now a multitude of signal molecules and the induced signal cascades have been identified, whereas this form of molecular regulation of signal transduction was defined as quorum sensing (QS). This term was given according to its special mechanism and was mentioned for the first time within a review of Fuqua et al. [15]. During the phase of cell growth a specific autoinducer molecule is produced as a function of increasing cell-population density, whereas the bacteria are also capable to detect and respond to the accumulation of these compounds. Due to this quorum sensing process the organisms can distinguish between high and low cell population density and can control gene expression in variation of cell population. Therefore bacteria have a special tool that allows a population of microorganisms to coordinately control the gene expression of the entire community [16]. Several organisms are capable to produce secretory
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enzymes that are capable of autoinducer degradation, while rendering the quorum sensing of competing bacteria mute and deaf. Therefore the development of novel antimicrobial therapeutics is currently focusing on quorum sensing signal molecules digesting enzymes. Such a mentioned cell-cell communication was firstly described by Hastings and Nealson [17,18], who elucidated the cell-density depending expression of luciferase of Vibrio fischeri. The microorganism emits light when habiting insight the specialized light organs of the host organism but not when free-living in the ocean. Up to now, several communication procedures were discovered, whereas distinct signal compounds were found depending on the species. Whereas gram-positive bacteria commonly utilize amino acids or small peptides for quorum sensing [19,20], derivates of fatty acids (acyl-homoserine lactones) serves as autoinducer compounds for gram-negative bacteria [21]. Furthermore, other systems concerning other principles than communication via lactones or amino acids and higher hierarchical systems for cross-species communication can be found elsewhere [21]. The discovery of these communication systems has an enormous impact on fermentation procedures of marine microorganisms. As already mentioned, most isolates stop production of metabolites and enzyme expression, respectively, when they are cultivated in traditional shake flasks. This effect results, because a well-agitated suspension culture does not reflect the natural growth conditions. By application of a modified roller-bottle bioreactor, which mimicked the intertidal environment Yan et al. could facilitate the production of antimicrobial compounds by two epiphytic isolates, which had stopped their production under planktonic growth conditions [22]. In addition it was already demonstrated that secondary metabolite production of marine microorganisms can be induced by cross-species interaction [23]. The authors applied a novel air-membrane surface bioreactor, which allows the bacteria to grow attached to a surface as a biofilm in contact with air. When the isolates get in contact with Bacillus sp. cultures the production of antimicrobial substances was induced, which suggests that there is a biofilm-specific cross-species signaling system. Since the organisms raise its enzyme/biocatalyst expression necessary for the production of the bioactive secondary metabolite in response to the quorum sensing stimuli, one can presumably transfer such approach also prior to enzyme recovery processes. Acylated homoserine lactones can be furthermore utilized as medium supplements within the cultivation process to improve growth of bacteria. For lake water bacteria Schink et al. reported that such a supplementation results in a significant improvement of growth performance [24]. Guan & Kamino described an increase of 3-8-fold of counting colony forming units by two marine derived samples, when cultivation occurs in the presence of acyl-homoserine lactones and commercial siderophore desferroixamine [25]. Recently, Lu et al. described the identification of the catalytic activity of a novel N-acylhomoserine lactonase from marine Bacillus cereus strain [26]. This development is quite interesting for pharmaceutical purposes, while AHL-lactonases are capable of disrupt the quorum sensing pathways in pathogenic organisms, wherefore these enzyme might be used within anti-infection therapeutic strategies. The responsible gene sequence of AHL-lactonase was cloned and expressed in E. coli, whereas the 28 kDa-protein was obtained with strong activity.
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3. Examples of Marine Enzymes As marine microorganisms are accessible with regard to cultivation, identification (by molecular phylogenetic methods) and bioprocessing, they are of major interest to researchers worldwide. The symbiotic nature (microorganisms found associated with various marine sponges, corals and with other species) and their occurrence in extreme environments (extremophiles) like hydrothermal vents have also been the areas of recent research. Within the following subchapters we are focusing therefore on interesting enzyme classes, applicable in research and industry.
3.1. Amylase and Pullulanase
The discovery of novel marine derived thermophilic enzymes capable to degrade starch will have a beneficial impact to industrial starch degrading processes. Such processes are carried out in large-scale dimensions for the production of sugars. During starch processing several successive steps, which involves different microbial enzymes are required [27]. The first step of the degradation is carried out at temperatures of about 95-105 °C at pH 6.0-6.5, whereas the subsequent step is done at pH 4.5 and 60 °C. The decline of the temperature is necessary, since more suitable enzymes are not available. While discovering novel thermophilic starch degrading enzymes such as α-amylase, pullulanase, glucoamylase and xylose isomerase a significant improvement of the process will be possible, due to the liquefaction and saccharification process might be achieved within one step under the same conditions [28]. Advantageous thermostable α-amylases and pullulanases showing a temperature optimum of more than 100 °C were found for example in the archaea Staphylothermus marinus, Pyrococcus furiosus, Pyrodictum abyssi [29]. Furthermore, amylases expressed by cold adapted, psychrophylic marine strains showing a well performance of starch digestion at low temperatures. Therefore an application as additive in detergents is highly recommended (see 3.2) [30]. Furthermore, pullulanase, glucosidase and amylase activities could be also detected in four archaeal strains related to the genus Thermococcus [31], whereas Brown and Kelly [32] purified extracellular pullulanases from cell free culture supernatants of the marine thermophilic archaea Thermococcus litoralis and Pyrococcus furiosus. Such enzymes from T. litoralis and P. furiosus appeared to represent highly thermostable amylopullulanases, versions of which however, have been isolated from less thermophilic organisms. Gantelet and Duchiron isolated the extremely thermophilic archaeon Thermococcus hydrothermalis from a deep sea hydrothermal vent in the East Pacific Rise which is capable to produce an extracellular pullulanase [33].
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3.2. Proteolytic Enzymes
Proteolytic enzymes, also named proteinases, are capable to degrade proteins to petides and amino acids. According to the world wide biocatalysts market, such proteinases represent the largest fraction. Especially serine alkaline proteases are used as additives for household laundering purposes, wherefore they have to resist inactivation by detergents and alkaline pH [28]. Furthermore, proteolytic enzymes are applied in leather processing, whereas proteinases with high keratinolytic and elastolytic properties are appreciated for soaking. Since proteinases can be also used within their reverse action mode, they are a valuable tool for the synthesis of peptides. Michels et al. [34] reported the properties of a hyperthermophilic and barophilic protease from Methanococcus jannaschii, which was the first protease to be isolated from a highpressure-high-temperature adapted organism. A Pyrococcus abyssi derived protease was isolated and purified from the supernatant of the hyperthermophilic archaebacteria by Dib et al. [35]. In addition a novel intracellular serine proteinase was identified from a marine aerobic hyperthermophilic archaeon Aeropyrum pernix [36]. That enzyme possesses a halflife of approximately 85 min at 100 °C and 12 min at 110 °C. While extensive investigations of shallow water and deep sea hydrothermal vents for the isolation of hyperthermophilic strains, mostly Pyrococcus sp. and Thermococcus sp., thermostable hydrolytic enzymes were characterized for potential applications. Vibrio species have been found to produce also a variety of extracellular proteases. Vibrio alginolyticus produces six proteases, including an unusual detergent resistant, alkaline serine exoprotease [37]. Shibata et al. discovered a novel metalloproteinase, named almelysin showing high activity at low temperatures and another proteinase from the culture supernatant of a marine Alteromonas sp. [38]. With respect to the fact that household washing is nowadays carried out under moderate temperatures, the demand for highly active cold-adapted enzymes proteinases is enormous. However, the production costs limit their application at present [30], but several patents have already been filed, which describe the utilization of proteinases from cold-adapted bacteria [39,40].
3.3. DNA-Polymerases
Enzymes capable of DNA processing are very important tools within contemporary molecular biology, since thermostable marine derived DNA polymerases play a major role in polymerase chain reaction and in a variety of other molecular biological applications such as DNA amplification, cloning, sequencing, or labeling [41]. Up to now several native and recombinantly expressed polymerases have been purified and characterized [42]. A second generation of thermostable polymerase chain reaction (PCR) enzymes has already been harvested from bacteria living near thermal vents on the ocean floor, and is marketed as Vent® and Deep Vent® Polymerases [43]. These enzymes feature a much better performance with respect to the error rate. Compared to the Taq polymerase the detectable error rate of these second generation polymerases is ten-fold lower [44].
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3.4. Halogenating Enzymes
Halogenated natural products have a widespread occurrence in natural environments. Up to now more than 4000 different organohalogens are known. Most of such compounds contain chlorine respectively bromine, whereas only minor amounts of fluoro- or iodohaloorganic compounds have been detected [45]. Due to the relatively high concentration of bromide found in sea water, bromination of aromatic and heteroaromatic molecules as well as bromination of isoprenoid metabolites can be found quite often in comparison to fresh water samples [46,47]. The natural product formation can be divided in abiotic respectively biogenous processes, whereas latter one require the availability of particular enzymes, which can be divided into two subclasses of halogenating enzymes: halogenases and haloperoxidases. In contrast to haloperoxidases, halogenases (flavin-dependent halogenases, mononuclear non-heme iron halogenases) are capable of catalyzing the regioselective formation of carbon halogen bonds and are therefore of peculiar interest for applications in White Biotechnology, since the commonly applied toxic halogenating agents could be substituted through the less harmful halides [48,49] and furthermore fewer by-products are produced [50]. Reaction of halogenases often occurs at positions 4, 5, 6 and 7 of tryptophanderived rings of natural products as well as halogenation of tyrosines at the ortho position and mono- and di-chlorination of pyrroles [51,52]. Due to their application as intermediates in Pd-catalysed carbon coupling reactions they are also of tremendous interest in the production of organic fine chemicals. While halogenated natural products often have a higher bioactivity than their unhalogenated counterparts [53], they will be quite prospective especially for pharmaceutical purposes. Most halogenation reactions are oxidative [54], but quite recently a novel non-enzymatic non-oxidative strategy was elucidated, which is responsible for halogenation of enediyne-derived macrolides in marine actinomycetes [55]. Within the frame of a “proof-of-concept-study” a Streptomyces sp. derived recombinantly expressed FADH2-dependent tryptophan-5-halogenase is currently under investigation, since the halogenated tryptophan may serve as a possible serotonin precursor for pharmaceutical applications [56]. While first experiments results only low enzyme activities, the authors applied a genetic algorithm to improve the enzyme’s performance and the conditions for the biotransformation process, respectively. Thus, the yield of the halogenated product was improved from 3.5 to 65%.
4. Molecular Biology Cloning ribosomal ribonucleic acid (rRNA) genes from mixed microbial assemblages is done to determine the phylogenetic identity of population constituents. Such cultivationindependent molecular phylogenetic surveys have revealed an astounding number of novel phylogenetic lineages [57]. Recent advances include the recovery of greater overall amounts of DNA in environmental DNA libraries. Rapid progress in high throughput screening, sequencing and robotics have also greatly facilitated a more thorough analysis of the recovered clones. These technological advances are vastly improving the economic and technical feasibility of cloning, screening and sequencing large numbers of clones derived
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from natural environments. There has been a good deal of interest in recovering microbial DNA from soil, with most studies concentrating on bioprospecting for drugs, enzymes and other natural products. This type of approach has been in use now for nearly a decade in the biotechnology industry [58]. Nowadays genetic engineers not only catalogue rRNAs (or other single genetic loci), but also determine large portions of the genomic content found within naturally occurring microbial communities. Different opportunities have become perceptible through this new approach. Bioprospecting, characterization of uncultivated microbes and microbial population genomics are advancing by its application [59]. In the early 1990s, molecular biology was fortified with the use of already mentioned thermostable DNA polymerases and the PCR [60] which became a major tool for phylogenetic diversity study of single genetic loci, especially rRNA genes [61]. Investigations in this sphere of research should have immense commercial applications. The cDNA encoding silicatein, the first silica synthesizing enzyme from the sponge Suberites domuncula was used as a probe to study the potential role of silicate on the expression of the silicatein gene. It was found that after increasing the concentration of soluble silicate in the seawater medium, this gene is strongly upregulated. This discovery has led to the European Community funded project to develop new routes for the structure-controlled biofabrication of silica nanostructure materials for biosensors, biomedical uses and bio-semiconductors by diatoms and siliceous sponges and for the industrial and medical application of the enzymes involved in synthesis and dissolution of biogenic silica.
5. Downstream Processes for Recovering Marine Enzymes While downstream processing is certainly one of the most underestimated steps in bioprocessing, this process step has the most significant effect on the complete process costs, especially for recovering and purification of pharmaceutical products. Interestingly research in this field is carried out only in a minor extent compared to those focusing on fermentation as well as up-stream techniques. A downstream process commonly starts from diluted raw material and tries to provide a highly concentrated, purified, and dried target compound. The target compound/enzyme can be provided by the expression system in extra- and intracellular manner. In the case of extra-cellular expression the target compound is available among other soluble (medium components, side products) and insoluble (particles, intact organisms) fractions. If the target is expressed intra-cellular the biomass matter must be disrupted prior to start the purification procedure. To obtain the unaltered native bioactive proteins, an enzymatic disintegration process by application of cell-wall digesting enzymes such as lysozyme is recommended. In general, all downstream processes try to remove the insoluble particles and afterwards the product is isolated. After a further purification step the process ends with the polishing [62]. A flow chart presenting the significant steps of such a process is given in Figure 1.
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Fermentation broth
Cell separation
Cell disintegration
Removal of Cell debris
Primary separation
Purification
Polishing
Packaging Figure 1. Flow chart of a general downstream process, according to [63].
The primary separation step comprises the removal of dissimilar compounds from the broth and accomplishing an essential volume reduction, whereas contemporary recovery procedures utilize such beneficial techniques like ion-exchange chromatography. This chromatographic procedure is often applied in order to remove either major contaminants from the broth or the target from the broth. Such chromatographic techniques – based on columns or membranes – are often an integral part of the purification step. Within the frame of the following subchapters several successful strategies for recovery of marine enzymes are described using such sophisticated techniques as membranes for ionexchange chromatography as well as affinity chromatography approaches. A deeper insight into basic concepts of more conventional separation techniques can be found elsewhere as well as further reading recommendations [63,64].
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5.1. Cell Disintegration
If the target enzyme is an intra-cellular expressed protein, which is not released to the fermentation broth during cultivation, the biomass must be disrupted to liberate the enzyme. Such a disintegration of biomass can be achieved by means of mechanical, chemical and enzymatic procedures. The method of choice is commonly the procedure which releases as much as possible of the product. But depending on the mechanism, parts of the target may suffer structure alteration and become inactive. Therefore very effective disruption and fast procedures are desired. Due to its short residence time, low operating costs, and contained operation mechanical methods are often preferred [65], whereas the most commonly applied techniques involve such apparatuses as homogenizers [66,67] or bead mills [67]. Homogenizers consist of a positive-displacement pump, which supplies the aqueous sample at high pressure through a tiny nozzle or an orifice valve device. Disruption occurs by shear forces and impingement of the cells onto the valve. The disruption rate depends on upstream pressure and geometry of the valve, whereas the operation is performed by multipass operation. However, temperature and velocity of the sample influences the performance of the disruption process. Thus, during cell disintegration heat is generated which may lead to denaturation of the target enzyme. With respect to its easy handling (sterilization, cleaning, marginal maintenance requirements) and its high degree of flexibility concerning the sample type this kind of homogenizer is been in use in several biotechnological laboratories. The also widely applied bead mills applies horizontal grinding chambers filled with small beads such as glass beads or more resistant zirconium oxide, zirconium silicate or titanium carbide and several others for disruption of (bio)materials [68]. Suspended biomass is introduced continuously into the grinding chamber. The turning speed of the grinding chamber has a tremendous influence concerning the disruption performance. During the grinding process the cells break up and release the target enzyme, whereas the resulting homogenate is separated from the grinding media by mechanical means. Unlike homogenizers a satisfying disruption level can be accomplished within a single run, whereupon a better performance of temperature distribution results. This effect allows additionally an improved temperature control. However, sterilization and cleaning is much more difficult and occurring bead abrasion leads to impurities of the homogenate. A phylogentic analysis of microbial communities present in marine sediments involving the usage of a bead mill is described by Gray and Herwig [69]. They report an operation protocol, which can be applied for efficient lysis of cells as well as recovery of DNA from marine sediments prior to phylogentic molecular studies, including 16S rRNA genes. Alternatively bead mills can be also operated in a batch mode, whereas the grinding chamber is not completely filled by the beads and agitated for reaching the liberation of intracellular compounds through the occurred shear stress. If the grinding chamber is used in such a mode the device is named mixer mill. Another method for accomplishing a convenient product release in a lab-scale from biomass utilizes ultrasound. By this technique the cells are disrupted by shear forces and cavitation, whereas cooling of the cell suspension is necessary to prevent heat denaturation. A cell lysis under mild conditions can be achieved by the use of hydrolyzing enzymes. Such enzymes degrade the organism’s cell wall and disintegration occurs as a result of
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osmotic pressure, which induces a blasting of the fermentative generated free protoplast. The lytic effects of enzymes are specific to particular groups of cell types and can be assigned to variations in cell wall composition. The most efficient bacterial cell wall digestion can be accomplished by lysozyme from hens’ egg origin. Due to the marginal equipment which is required for such a cell disintegration the process can be easily arranged, wherefore enzymatic lysis is also an option for large scale processes [70]. It is well known, that lysozyme from hens’ egg has also an antitumor function [71], research is focusing also on novel marine-derived lysozyme as possible anti-cancer agents [72]. In addition it could be shown that cold-adapted lysozyme extracted from marine bacterium possess some beneficial features in comparison to the chicken-derived enzyme such as broad range of pH optimum and temperature optimum, and more broad spectrum concerning antibiotic properties [73,74]. A more sophisticated method for sample disruption is provided by electrical pulse techniques. This approach is based on electro-induced changes in the cell envelope leading to a release of intracellular proteins without the formation of debris and permits the treatment of large volumes. For example, while treatment of yeasts by a couple of electric field pulses almost 90% of the total activity can be obtained without any further nor previous treatment of the biomass, whereas handling of at least 20% wet weight suspensions is feasible [75].
5.2. Membrane Adsorbers in Downstream-processing
The utilization of membrane adsorbers for the isolation of marine enzymes has several advantages compared to conventional column chromatography (e.g. lower manufacturing costs, no diffusion-controlled exchange kinetics, easier handling and up-caling) leading to a better process performance [76]. The fundamental concept of membrane adsorbers is to raise the separation efficiency by maximizing mass transfer, while modified microporous membranes serve as stationary matrix. Such membranes can be converted into efficient adsorbers by attaching functional groups to the inner surface of the synthetic membrane carrier. Several distinct chromatographic techniques such as ion-exchange, affinity adsorption or immobilized metal affinity chromatography can be carried out by these membranes. Commercial available are membrane ion exchangers of strong acidic (sulfonic acid), strongly basic (quarternary ammonium), weakly acid (carboxylic acid), and weakly basic (diethylamine), as well as chelating membranes functionalized with iminodiacetate (IDA) applicable for immobilized metal affinity chromatography. As already mentioned the membrane adsorber technology has several significant advantages compared to conventional chromatographic techniques. Due to its structure the membranes are capable of binding proteins without a limiting diffusion process, wherefore loading and elution can be performed at high fluxes which allows beneficial short cycle times. Unlike traditional chromatography the compressibility of the membrane can be neglected under moderate process control. Additionally, channelling does not occur and the pressure distribution inside a module is designed to perform plug flow through the module. Owing to this features sharp breakthrough curves results by application of membrane adsorbers. With regard to its modular concept the downstream process is easy to scale-up. Concerning the high levels of hygienic standards of several process industries
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(e.g. beverage, dairy, pharmaceuticals, cosmetics) the membrane units are accessible for (CIP (cleaning in place) and the process validation is commonly simplified due to introduction of standard products and validation service of suppliers. Currently commercial available are membranes for laboratory as well as process scale, whereas the modules and systems can be adopted to the special needs of the specific separation process to achieve optimal performance. Characteristic applications for membrane adsorber technology can be summarized as concentration of minor proteins [77] and monoclonal antibodies [78], removal of contaminants (e.g. DNA, endotoxins) [79] and reduction of virus content [80]. The application of membrane adsorbers for recovery of a marine sulfite-oxidizing enzyme was successfully established [77]. The enzyme (sulfite oxidase; sulfite: acceptor oxidoreductase) catalyzes the oxidation of sulfite to sulfate. In mammalian tissues the physiological importance of sulfite oxidase is due to its role as a terminal enzyme in the degradation of sulfur containing amino acids. Furthermore, it is important in the detoxification of endogenous sulfite and sulfur dioxide. Such kind of enzyme has been located in several mammalian tissues, such as the liver, but also in plants and bacteria. The marine sulfite-oxidizing enzyme is produced by Sulfitobacter pontiacus, a Gram-negative bacterium that was isolated from water samples taken from a depth of 100-140 m at the H2SO2 interface in the eastern part of the Black Sea [81]. The strain is strict heterotrophic and is unable to grow autotrophically on H2, thiosulfate or sulfite. The organism is strict aerobe and requires NaCl (5-80 g/L, optimum 20-25 g/L). Temperature and pH ranges are 4-35°C and pH 6.5-8.5. While studying the metabolism of the microorganism Sorokin et al. found that in acetate-limited continuous culture, the bacterium tolerates after adaptation such high sulfite concentration of up to 63 mM [82]. In addition they discovered that oxidation of sulfite to sulfate, accomplished by a highly active AMP-independent soluble sulfite-oxidizing enzyme, leads to an increase in biomass concentration. This result indicates that the strain is capable to utilize sulfite as an additional source of energy. Ulber et al. are focusing on recovery of the sulfite-oxidizing enzyme from S. pontiacus on a larger-scale, while establishing also a fedbatch operation mode for production of the sulfite-utilizing enzyme [77,83]. After the cultivation the enzyme was isolated as follows: • • • • •
Centrifugation of the fermentation broth Ultrasonication of re-suspended biomass Centrifugation of cell debris Cation exchange of the supernatant Ultrafiltration of the obtained active fractions
Within this purification concept the application of the cationic membrane adsorber is an integral part of the downstream processing. Figure 2 illustrates a typical elution profile. While implementing the membrane adsorber into the downstream process as mentioned above, a highly purified sulfite-oxidizing enzyme is obtained from the marine bacterium Sulfitobacter pontiacus. The resulting activity of the enzyme in comparison with commercially available sulfite oxidase from chicken liver origin was approximately 25-fold higher.
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0,60
0,40 1,2
conductivity UV-absorption enzyme activity
0,35 1,0
0,45
0,8
0,40 0,6 0,35 0,30
0,4
active
0,30
enzyme activity [U/ml]
rel. absorption280nm
0,50
0,25 0,20 0,15 0,10
rel. conductivity
0,55
fractions
0,25
0,05
0,2
0,20
0,00 0,0 0
20
40
60
80
Time [min]
Figure 2. Elution profile of cation exchange chromatography of crude cell extract of Sulfitobacter pontiacus for recovery of sulfite-oxidizing enzyme using membrane adsorbers (Sartorius, Germany); the conditions were adjusted as follows: 0-5 min buffer A (20 mM sodium acetate buffer, pH 4.6), 5-20 min 0-20% buffer B (20 mM sodium acetate buffer, 0.2 M NaCl, pH 4.6), 20-60 min 20-35% buffer B, 60-70 min 35-100% buffer B, 70-80 min buffer B; flow rate: 2 ml/min [83].
Table 2. Purification of a sulfite-oxidizing enzyme from Sulfitobacter pontiacus [77] Fraction
Vol.
Vol. activity
Protein
Total protein
Spec. activity
Units
Yield
[mL]
[U/mL]
[µg/mL]
[µg]
[U/mg]
[U]
[%]
Crude extract
10
3.17
141
1415
22.5
31.76
100
IEC-membrane
24
0.61
3.42
82.1
178.0
14.61
46
UF
2.2
6.24
13.57
29.9
460
13.73
43
An overview of the resulting concentration factors as well as the yield of each purification step is given in Table 2.
5.3. Affinity Adsorption (Chromatography)
Affinity adsorption relies on highly specific binding interactions, unlike conventional separation techniques which rely mainly on molecular size, charge or solubility. Due to this high specific interaction of the stationary system of the chromatographic column, purification factors of almost 1000 and high recovery rates can be accomplished. Up to know this
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technique is well established for several downstream processes [84,85]. The process performance results from the formation of a reversible complex between the target (protein) and the ligand, whereas the ligand is commonly applied in an immobilized form (insoluble matrix). Affinity adsorption techniques are already introduced into downstream processes of enzymes and antibodies from marine origin. For instance Berteau et al. describe the purification and characterization of a marine derived alpha-L-fucosidase within a three step chromatographic process including an essential affinity chromatographic step based on the glycosidase inhibitor analogue 6-amino-deoxymannojirimycin as ligand [86]. Such fucosidases are capable to degrade high molecular sulfated fucoidans without loss of sulfate nor structure alteration into so-called sulfated low molecular fucoidans, whereas the biological activity is increased. Such smaller subunits of the fucoidan polymer are of tremendous interest especially for pharmaceutical purposes since these subunits show a similar efficacy as animal derived heparin [87,88] and the sulfated fragments possess also anti-viral [89,90] and anti-tumor activity [91], respectively. Especially owing to its heparinlike behaviour, fucoidan preparations have been proposed as alternatives to the anticoagulant heparin, prepared from mammalian mucosa. Therefore the application of such pharmaceutically useful compounds of vegetable origin is less likely to entail infectious agents (e.g. viruses or prions) [92]. A sophisticated approach for isolation and characterization of the blood group B specific lectin from the red marine alga Ptilota plumose was carried out by Sampaio et al. [93]. After loading the aqueous extract onto a Sephadex G-200 column (1.6 × 18 cm), equilibration, and elution with PBS containing 1 mM CaCl2 at 10 mL/h (until the absorbance of the effluent is less than 0.05) the adsorbed proteins were eluted with 50 mL 0.1 M D-glucose in PBS containing 1 mM CaCl2. Thus, the authors obtained a concentration factor of 212, while the specific activity increased from 218 U/mg in the crude extract to 46,282 U/mg in the processed purified material. Haukson et al. described the recovery of a particularly heat-labile alkaline phosphatase from marine microorganism origin, using a gentle single-step affinity chromatography procedure on agarose-linked L-histidyldiazobenzylphosphonic acid, which results 54% yield by a concentration factor of 151 [94]. Moreover, other enzymes from marine organisms origin were already purified by application of affinity chromatographic techniques such as serine proteases [95], thermostable phosphatase [96], sialyltransferase [97] or exopolyphosphatases [98].
6. Summary During the past decade, a couple of novel marine enzymes from exotic sources with characteristic and beneficial traits have reached the stage of commercial production. The main problem is often the limited availability of the enzymes, considering enzyme expression under laboratory-scale conditions as well as their commonly low stability, which hampers their utilization for industrial scale processes. As mentioned in chapter 2, several research is done within the field of bioreactor design. While immobilized cell technology has received also attention by marine researchers, one can expect novel industrial applications of stabilized enzymes for operation under psychrophilic, mesophilic as well as thermophilic
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conditions. However, psychrophilic and inducible expression systems were developed recently. With regard to basic molecular research in the field of molecular biology this will support a beneficial expression under very mild conditions, without significant activity loss [99]. Due to the fact, that up to now only a minor amount of the marine diversity was screened with respect to novel organisms as well as enzymes, one can expect a prospective future within this emerging field.
7. References [1] [2] [3] [4]
[5] [6] [7]
[8] [9] [10]
[11]
[12] [13] [14]
Watve, M; Tickoo, R; Jog, M; Bhole, B. How many antibiotics are produced by the genus Streptomyces? Arch. Microbiol., 176 (2001), 386-390. Fenical, W. Chemical Studies of Marine Bacteria: Developing a new Resource. Chem. Rev., 93 (1993), 1673-1683. Faulkner, DJ. Marine Natural Products. Nat. Prod. Repts., 18 (2001), 1-49. Wagner-Döbler, I; Beil, W; Lang, S; Meiners, M; Laatsch, H. (2002). Integrated approach to explore the potential of marine microorganisms for the production of bioactive metabolites. In T. Scheper (Ed.), Advances in Biochemical Engineering/Biotechnology, Vol. 74, 207-238. Berlin, Heidelberg, New York, Springer Verlag. Proksch, P; Edrada, RA; Ebel, R. Drugs from the seas - current status and microbiological implications. Appl. Microbiol. Biotechnol., 59 (2002), 125-134. Holzgrabe, A; Bechthold, A. Paradigmenwechsel in der Entwicklung antiinfektiver Chemotherapeutika. Chemotherapie Journal, 8 (1999), 69-78. Debashish, G; Malay, S; Barindra, S; Joydeep, M. (2005). Marine Enzymes. In Y. Le Gal & R. Ulber (Eds.), Advances in Biochemical Engineering/Biotechnology, Vol. 96, 189-218. Austin, B. (1988). Marine Microbiology. Cambridge University Press, New York. Flemming, HC; Wingender, J. Biofilme - die bevorzugte Lebensform der Bakterien. Biol. Unserer Zeit 3 (2001), 169-180. Button, DK; Schut, F; Quang, P; Martin, R; Robertson, BR. Viability and isolation of marine bacteria by dilution culture: Theory, procedures, and initial results. Appl. Environ. Microbiol., 59 (1993), 881-891. Gledhill, M; van der Berg, CMG;. Determination of complexation of iron (III) with natural organic complexing ligands in seawater using cathodic stripping voltammetry. Mar. Chem., 47 (1994), 41-54. Tortell, PD; Maldonado, MT; Granger, J; Price, NM. Marine bacteria and biogeochemical cycling of iron in the oceans. Nature, 383 (1996), 330-332. Neilands, JB; Siderophores: Structure and function of microbial iron transportcompounds. J. Biol. Chem., 270 (1995), 26723-26726. Stock, AM; Robinson, VL; Goudreau, PN. Two-component signal transduction. Annu. Rev. Biochem., 69 (2000), 183-215.
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[15] Fuqua, WC; Winans, SC; Greenberg, EP. Quorum sensing in bacteria: the LuxR-LuxI family of cell density-responsive transcriptional regulators. J. Bacteriol., 176 (1994), 296-275. [16] Schauder, S; Bassler, BL. The language of bacteria. Genes & Development, 16 (2001), 1468-1480. [17] Hastings, JW; Nealson, KH. Bacterial biolumiscence. Annu. Rev. Microbiol., 31 (1977), 549-595. [18] Nealson, KH; Hastings, JW. Bacterial bioluminescence: Ist control and ecological significance. Microbiol. Rev., 43 (1979), 496-518. [19] Kleerebezem, M; Quadri, LEN; Kuipers, OP; de Vos, WM. Quorum sensing by peptide pheromones and two-component signal transduction systems in Gram-positive bacteria. Mol. Microbiol., 24 (1997), 895-904. [20] Lazazzera, BA; Grossman, AD. The ins and outs of peptide signalling. Trends Microbiol., 7 (1998), 288-294. [21] Whitehead, NA; Barnard, AML; Slater, H; Simpson, NJL; Salmond, GPC. Quorumsensing in Gram-negative bacteria. FEMS Microbiol., Rev. 25 (2001), 365-404. [22] Yan, L; Boyd, KG; Burgess, JG. Surface attachment induced production of antimicrobial compounds by marine epiphytic bacteria using modified roller bottle cultivation. Mar. Biotechnol., 4 (2002), 356-366. [23] Yan, L; Boyd, KG; Adams, DR; Burgess, JG. Biofilm-specific cross-species induction of antimicrobial compounds in bacilli. Appl. Environ. Microbiol., 69 (2003). [24] Bussmann, I; Philipp, B; Schink; B. Factors influencing the cultivability of lake water bacteria. J. Microbial Met., 47 (2001) 41-50. [25] Guan, LL; Kamino, K. Bacterial response to siderophore and quorum-Sensing chemical signals in the seawater microbial community. BMC Microbiol., 1 (2001), 27. [26] Lu, X; Yuan, Y; Xue, XL; Zhang, GP, Zhou, SN. Identification of the critical role of Tyr-194 in the catalytic activity of a novel N-acyl-homoserine lactonase from marine Bacillus cereus strain Y2. Curr. Microbiol., 53 (2006), 346-350. [27] Antranikian, G. (1992). Microbial degradation of starch. In G. Winkelmann(Ed.), Microbial degradation of natural products. VCH, Weinheim, 27-56. [28] Ladenstein, R; Antranikian, G. (1998). Proteins from Hyperthermophiles: Stability and enzymatic catalysis close to the boiling point of water. In G. Antranikian (Eds.), Advances in Biochemical Engineering/Biotechnology, Vol. 61, 37-85. [29] Sunna, A; Moracci, M; Rossi, M; Antranikian, G. Glycosyl hydrolases from hyperthermophiles. Extremophiles, 1 (1997), 2-13. [30] Marx, JC; Collins, T; D’Amico, S; Feller, G; Gerday, C. Cold-adapted enzymes from marine antarctic microorgansims. Mar. Biotechnol., 9 (2007), 293-304. [31] Legin, E; Ladrat, C; Godfroy, A; Barbier, G. Thermostable amylolytic enzymes of thermophilic microorganisms from deep-sea hydrothermal vents. Comptes Rendus de l'Académie des Sciences - Series III - Sciences de la Vie, 320 (1997), 893-898
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[32] Brown, SH; Kelly, RM. Characterization of Amylolytic Enzymes, Having Both alpha1,4 and alpha-1,6 Hydrolytic Activity, from the Thermophilic Archaea Pyrococcus furiosus and Thermococcus litoralis. Appl. Environ. Microbiol., 59 (1993), 2614-2621. [33] Gantelet, H; Duchiron, F. Purification and properties of a thermoactive and thermostable pullulanase from Thermococcus hydrothermalis, a hyperthermophilic archaeon isolated from a deep-sea hydrothermal vent. Appl. Microbiol. Biotechnol. 49 (1998), 770-777. [34] Michels, PC; Clark, DS. Pressure-enhanced activity and stability of a hyperthermophilic protease from a deep-sea methanogen. Appl. Environ. Microbiol. 63 (1997), 3985-3991. [35] Dib, R; Chobert, JM; Dalgalarrondo, M; Barbier, G; Haertle, T. Purification, molecular properties and specificity of a thermoactive and thermostable proteinase from Pyrococcus abyssi, strain st 549, hyperthermophilic archaea from deep-sea hydrothermal ecosystem. FEBS Lett., 431 (1998), 279-284 [36] Chavez Crooker P; Sako, Y; Uchida, A. Purification and characterization of an intracellular heat-stable proteinase (pernilase) from the marine hyperthermophilic archaeon Aeropyrum pernix K1. Extremophiles 3 (1999), 3-9. [37] Biotechnology for the 21st century: new horizons 5. Opportunities in marine biotechnology and aquaculture. (1995). Reports prepared by Biotechnology Research Subcommittee, Committee on Fundamental Science and National Science and Technology Council, USA. [38] Shibata, M; Takahashi, S; Sato, R; Oda, K. A novel metalloproteinase, almelysin, from a marine bacterium, Alteromonas sp. No. 3696: purification and characterization. Biosci. Biotechnol. Biochem. 61 (1997), 710-715. [39] Quamrul, H; Eiichi, T: Cold-active protease CP70. Patent 1998, No. US6200793. [40] Hiromi, K; Mikio, T; Naoko, F; Susumu, I; Toru, K; Yasuhiko, O; Katsuhisa, S; Mitsuyoshi, O. Cold alkaline protease, microorganism producing the same, process for producing the same, and detergent compositions and food processing enzyme preparations containing the same. Patent application 1996, No. WO9743406. [41] Alguéres, SMC; Almeida, RV; Clementino, MM; Vieira, RP; Almeida, WI; Cardoso, AM; Martins, OB. Exploring the biotechnological applications in the archaeal domain. Brazilian J. Microbiol., 38 (2007), 398-405. [42] Perler, FB; Kumar, S; Kong, H. Thermostable DNA polymerases. Adv. Protein Chem., 48 (1996), 377-435. [43] Grace, ES. (1997). Biotechnology in Seas and Trees. In Biotechnology Unzipped: Promises and Realities. Natl Academies Press, Washington D.C. [44] Southworth, MW; Kong, H; Kucera, RB; Ware, J; Jannasch, HW; Perler, FB. Cloning of thermostable DNA polymerases from hyperthermophilic marine Archaea with emphasis on Thermococcus sp. 9 degrees N-7 and mutations affecting 3’-5’exonuclease activity. Proc. Natl. Acad. Sci. USA, 93 (1996), 5281-5285. [45] Gribble, GW. Amazing organohalogens. Am. Sci. 92 (2004), 342-349. [46] Butler, A; Walker, JV. Marine haloperoxidases. Chem. Rev., 93 (1993), 1937-1944.
Marine Enzymes for Biocatalysis – Production, Isolation and Applications
277
[47] Carter-Franklin, JN; Butler, A. Vanadium bromoperoxidase-catalyzed biosynthesis of halogenated marine natural products. J. Am. Chem. Soc., 126 (1996), 15060-15066. [48] Hammer, PE; Hill, DS; Lam, ST; van Pée, KH; Ligon, JM. Four genes from Pseudomonas fluorescens that encode the biosynthesis of pyrrolnitrin. Appl. Environ. Microbiol., 63 (1997), 2147-2154. [49] Keller, S; Wage, T; Hohaus, K; Hölzer, M; Eichhorn, E; van Pée, KH. Purification and Partial Characterization of Tryptophan 7-Halogenase (PrnA) from Pseudomonas fluorescens. Angew. Chem. Int. Ed., 39 (2000), 2300-2302. [50] Hasegawa, M; Yamada, K; Nagahama, Y; Somei, M. The chemistry of indoles Part 94 - A novel methodology for preparing 5-chloro- and 5-bromotryptamines and tryptophans, and its application to the synthesis of (+/-)-bromochelonin B. Heterocycles, 51 (1999), 2815-2821. [51] van Pée, KH. Biosynthesis of halogenated metabolites by bacteria. Annu. Rev. Microbiol., 50 (1996), 375-399. [52] van Pée, KH. Microbial biosynthesis of halometabolites. Arch. Microbiol., 175 (2001), 250-258. [53] Naumann, K; Chlorchemie der Natur. Chem. Unserer Zeit; 27 (1993); 33–41. [54] Fujimori, DG; Walsh, CT. What’s new in enzymatic halogenations. Curr. Opin. Chem. Biol., 11 (2007, 553-560. [55] Perrin, CL; Rodgers, BL; O’Connor, JM. Nucleophilic addition to a p-benzyne derived from an enediyne: a new mechanism for halide incorporation into biomolecules. J. Am. Chem. Soc., 129 (2007), 4795-4799. [56] Muffler, K; Retzlaff, M; van Pée, KH; Ulber, R. Optimisation of halogenase enzyme activity by application of a genetic algorithm. J. Biotechnol., 127 (2007); 425-433. [57] Pace, NR. A molecular view of microbial diversity and the biosphere. Science, 276 (1997), 734-740. [58] Short, JM. Recombinant approaches for accessing biodiversity. Nat. Biotechnol., 15 (1997), 1322-1323. [59] DeLong, EF. Microbial population genomics and ecology. Curr. Opin. Microbiol., 5 (2002), 520-524. [60] Saiki, RK; Gelfand, DH; Stoffel, S; Scharf, SJ; Higuchi, R; Horn, GT; Mullis, KB; Ehrlich, HA. Primer-directed enzymatic amplification of DNA with a thermostable DNA-polymerase. Science, 239 (1988), 487-491. [61] Giovannoni, SJ; Britschgi, TB; Moyer, CL; Field, KG. Genetic diversity in sargasso sea bacterioplankton. Nature, 345 (1990), 60-63. [62] Belter, PA; Cussler, EL; Hu W-S. (1988). Bioseparations. Wiley, New York. [63] Muffler, K; Ulber, R. (2005). Downstream processing in marine biotechnology. In Y. Le Gal & R. Ulber (Eds.), Advances in Biochemical Engineering/Biotechnology, Vol. 97, 64-103. [64] Becker, T; Breithaupt,D; Doelle, HW; Fiechter, A; van Griensven, M; Kasper, C; Lütz, S; Pörtner, R; Schlegel, HG; Sell, D; Shimizu, S; Stahl, F; Suck, K; Ulber, R;
278
[65] [66] [67] [68] [69] [70] [71] [72]
[73]
[74] [75] [76]
[77]
[78] [79] [80]
[81]
K. Muffler, J. Mukherjee and R. Ulber
Wegener, J; Würges, K; Yamada, H; Zorn, H. (2007). Biotechnology, in Ullmann’s Encyclopedia of Industrial Chemistry, Electronic Release, Wiley-VCH, Weinheim. Hatti-Kaul, R; Mattiasson, B. (2003). Isolation and purification of proteins. Marcel Dekker, New York. Asenjo, JA. (1990). Separation processes in Biotechnology. Marcel Dekker, New York. Goldberg, E. (1997). Downstream processing. Chapman & Hall, London. Middelberg, APJ. Process-scale disruption of microorganisms. Biotech. Adv., 13 (1995), 491-551. Gray, JP; Herwig, RP. (1996). Phylogenetic analysis of the bacterial communities in marine sediments. Appl. Environ. Microb., 62, 4049-4059. Stephanopoulos, G. (1993). Biotechnology. VCH, New York. Sava, G; Benetti, A.; Ceschia, V; Pacor, S. Lysozyme and cancer: role of exogenous lysozyme as anticancer agent. Anticancer. Res., 9 (1989), 583-591. Ye, JL; Wang, CB; Chen, XH; Guo, SB; Sun, M. Marine lysozyme from a marine bacterium that inhibits angiogenesis and tumor growth. Appl. Microbiol. Biotechnol., 77 (2008), 1261-1267. Wang, YJ; Sun, M; Zhang, YB; Hong, YG; Hao, JH; Liu, XP; Wang, CB. Studies on preparation and characteristic of the marine low temperature lysozyme. Mar. Fish. Res. 21 (2000), 54-63. Zou, YL; Sun, M; Wang, YJ. Purification and Characterization of a lysozyme from a marine microorganism. Chin. J. Biotech., 21 (2005) 420-424. Ganeva, V; Galutzov, B; Teissie, J. High yield electroextraction of proteins from yeast by a flow process. Anal. Biochem., 315 (2003), 77-84 Ulber, R; Plate, K; Reif, OW; Melzner, D. (2003). Membranes for protein isolation and purification. In R. Hatti-Kaul & B. Mattiasson (Eds.), Isolation and purification of proteins. Marcel Dekker, New York, 191-224. Ulber, R; Arndt, M; Muffler, K; Laroche, M. (2003). Enzymes from marine microorganisms. In Marine Biotechnology an overview of leading fields. Ifremer, Nantes, 9-16. Zhou, JX; Tressel, T. Basic concepts in Q membrane chromatography for large-scale antibody production. Biotechnol. Prog., 22 (2006), 341-349. Bell, CM; Guo, K; Wendel, HP. Endotoxin removal from albumin and saline solutions. Int. J. Artif. Organs. 30 (2007), 589-593. Kalbfuss, B; Wolff, M; Geisler, L; Tappe, A; Wickramasinghe, R; Thom, V; Reichl, U. Direct capture of influenza A virus from cell culture supernatant with Sartobind anionexchange membrane adsorbers. J. Membr. Sci., 299 (2007) 251-260. Sorokin, DY. Sulfitobacter pontiacus gen. nov., sp. nov. – A new Heterotrophic Bacterium from the Black Sea, Specialized of Sulfite Oxidation. Microbiology, 64 (1995), 295-305.
Marine Enzymes for Biocatalysis – Production, Isolation and Applications
279
[82] Sorokin, DY; Vedenina, IY; Grabovich, MY. Influence of sulfite on acetate metabolism in Sulfitobacter pontiacus growing in continuous culture. Microbiology, 68 (1999), 14-20. [83] Muffler, K; Ulber, R. Fed-batch cultivation of the marine bacterium Sulfitobacter pontiacus using immobilized substrate and purification of sulfite oxidase by application of membrane adsorber technology. Biotechnol. Bioeng., 99 (2008), 870875. [84] Roy, I; Gupta, MN. (2003). Selectivity in affinity chromatography. In R. Hatti-Kaul & B. Mattiasson (Eds.), Isolation and purification of proteins. Marcel Dekker, New York, 57-94. [85] Odde, DJ. (1997). Affinity adsorption. In E. Goldberg (Ed.), Handbook of downstream processing. Chapmann & Hall, London, 70-89. [86] Berteau, O; McCort, I; Goasdoue, N; Tissot, B; Daniel, R. Characterization of a new alpha-L-fucosidase isolated from the marine mollusk Pecten maximus that catalyzes the hydrolysis of alpha-L-fucose from algal fucoidan (Ascophyllum nodosum). Glycobiology, 12 (2002), 273-282. [87] Church, FC; Pratt, CW; Noyes, CM; Kalayanamit, T; Sherrill, GB; Tobin, RB; Meade, JB. Structural and functional properties of human alpha-thrombin, phosphopyridoxylated alpha-thrombin, and gamma-T-thrombin - Identification of lysyl residues in alpha-thrombin that are critical for Heparin and fibrin(ogen) interactions. J. Biol. Chem., 264 (1989), 18419-18425. [88] Nishino, T; Aizu, Y; Nagumo, T. The influence of sulfate content and molecularweight of a fucan sulfate from the brown seaweed ecklonia-kurome on ist antithrombin activity. Thromb. Res., 64 (1991), 723-731. [89] McClure, MO; Moore, JP; Blanc, DF; Scotting, P; Cook, GM, Keynes, RJ; Weber, JN; Davies, D; Weiss, RA. Investigations into the mechanism by which sulfated polysaccharides inhibit HIV-infection invitro. Aids Res. Hum. Retroviruses, 8 (1992), 19-26. [90] Baba, M; Snoeck, R; Pauwels, R; de Clerk, E. Sulfated polysaccharides are potent and selective inhibitors of various enveloped viruses, including herpes-simplex virus, cytomegalo-virus, vesicular stomatitis-virus, and human immunodeficiency virus. Antimicrob. Agents Ch., 32 (1988), 1742-1745. [91] Riou, D; Colliec-Jouault, S; DuSel, DP; Bosch, S; Siavoshian, S; LeBert, V; Tomasoni, C; Sinquin, C; Durand, P; Roussakis, C. Antitumor and antiproliferative effects of a fucan extracted from Ascophyllum nodosum against a non-small-cell bronchopulmonary carcinoma line. Anticancer Res., 16 (1996), 1213-1218. [92] Berteau, O; Mulloy, B. Sulfated fucans, fresh perspectives: structures, functions, and biological properties of sulfated fucans and an overview of enzymes active toward this class of polysaccharide. Glycobiology, 13 (2003), 29-40. [93] Sampaio, AH; Rogers, DJ; Barwell, CJ; Saker-Sampaio, S; Nascimento, KS; Nagano, CS; Farias, WRL. New affinity procedure for the isolation and further characterization
280
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[96]
[97]
[98]
[99]
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of the blood group B specific lectin from the red marine alga Ptilota plumosa. J. Appl. Phycol., 14 (2002), 489-495. Hauksson, JB; Andresson, OS; Asgeirsson, B. Heat-labile bacterial alkaline phosphatase from a marine Vibrio sp. Enzyme Microb. Technol., 27 (2000), 66-73. Croocker, PC; Sako, Y; Uchida, A. Purification and characterization of an intracellular heat-stable proteinase (pernilase) from the marine hyperthermophilic archaeon Aeropyrum pernix K1. Extremophiles, 3 (1999), 3-9. Bannikova, GE; Varlamov, VP; Miroshnichenko, ML; Bonch-Osmolovskaya, EA. Isolation of thermostable phosphatase from the hyperthermophilic archaeon Thermococcus pacificus by immobilized metal affinity chromatography. Mol. Biol. Int., 44 (1998), 363-370. Yamamoto, T; Nakashizuka, M; Kodama, H; Kajihara, Y; Terada, I. Purification and characterization of a marine bacterial beta-galactoside alpha 2,6-sialyltransferase from Photobacterium damsela JT0160. J. Biochem., 120 (1996), 104-110. Lorenz, B; Batel, R; Bachinski, N; Müller, WEG; Schröder, HC. Purification and characterization of 2 exopolyphosphatases from the marine sponge Tethya lyncurium. Biochim. Biophys. Acta, 1245 (1995), 17-28. Duilio, A; Tutino, ML; Marino, G. Recombinant protein production in Antarctic Gramnegative bacteria. Meth. Mol. Biol., 267 (2004), 225-237.
In: Biocatalysis Research Progress Editors: F. H. Romano, A. Russo
ISBN: 978-1-60456-619-2 © 2008 Nova Science Publishers, Inc.
Chapter XII
Immobilized Microbial Cells Applications and Mass Transfer Phenomena Venko Beschkov* Institute of Chemical Engineering, Bulgarian Academy of Sciences, 1113 Sofia, Bulgaria
Abstract Immobilized cells performance has been extensively studied in the last three decades. The advantages of this type of biocatalysts are in their multiple use, continuous operation and easy removal from the reaction mixture. Two main types of immobilization techniques exist: cell entrapment in gels and cell fixation on solid supports. The drawbacks of cell entrapment are in the diffusion limitations at substrate supply to and product removal from the gel particles. Such limitations do not exist in cases of cell fixation on solid supports but other transport phenomena arise, associated with the cell accumulation and concentration within small area. Both techniques admit changes in the cell physiology including microbial growth different from that in free culture. That is why additional impact on the net biocatalyst performance may cause the microbial growth in immobilized state and the possible cell leakage into the fermentation broth and their consecutive growth in a free state. The choice of immobilization method depends on the kinetics of the very microbial process, on the microbial growth, on the products of reactions, being possible inhibitors or catalysts, etc. The purpose of this article is to review and to discuss the advantages and the drawbacks of these two groups of immobilized biocatalysts from the point of view of the specific reaction kinetics and the mass transfer limitations. Mathematical modeling is employed to evaluate the effects of cell growth and detachment from the particles and *
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[email protected]
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their contribution in free and immobilized state. Experimental data are presented to illustrate the discussed effects.
Notations A a Bi C D k kL KM’ r R Rd t T = R2/D V X
particle area, m2; specific particle area, related to their volume, m-1; Biot number, (-); solute concentrations, kg m-3; diffusivity, m2 s-1; rate constant of first-order reaction, s-1; external mass transfer coefficient, m s-1; Michaelis constant, kg m-3; radial coordinate, m; particle radius, m; ratio of product to substrate diffusivity, (-); time, s; dimensionless time; reaction rate, kg m-3 s-1; biomass concentration, kg m-3;
Greek symbols
β
leakage factor, m;
β = β/R
dimensionless leakage factor, related to the particle radius;
ξ η μ
reaction efficiency, Eq. (4); degree of conversion, Eq. (7); microbial specific growth rate, s-1;
Φ S2
Thiele modulus, (-).
I. Introduction The research on the properties of immobilized microbial cells is pretty widespread. The interest toward this type of biocatalyst is provoked by different reasons. First of all, it is their multiple use and the chance to manage industrial microbial processes in a continuous way. The second one is the possibility to attain much higher cell concentrations compared to free cultures. The immobilization of whole cells is preferred to the immobilization of enzymes, particularly for large scale industrial applications. Preparation of these biocatalysts is cheaper
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because of the cheaper downstream processing: there is no use of enzyme concentration, isolation and purification. Along with the advantages, the applications of immobilized microbial cells have some peculiarities compared to the free ones and raise some problems. The immobilized cells may behave in a different way compared to the free ones. For example, optimum pH-shift was observed (Chibata et al. [1]). Next, the growth of immobilized cells might be different from the free ones. Furthermore, in general the immobilized cells may have different productivity, i.e. different yield coefficients. Another feature of immobilized cells is the possible growth and subsequent detachment of cells from the carrier (or polymer matrix) into the fermentation liquid. Therefore, the immobilized cells may serve as a biocatalyst and as cell donor as well. The most important drawback to the broader application of immobilized cells is the mass transfer resistance for the substrate penetration and the removal of products from the biocatalyst particles, particularly when the product is an inhibitor. In the present review these features of immobilized microbial cells are discussed. An attention is paid to the mathematical modeling and its use for quantitative estimation of different factors, like the cell detachment rate, the contribution of free and immobilized cells for the overall fermentation process, the importance of the cell profile in the matrix or in the biofilm, etc.
II. Immobilization Methods There are two main groups of immobilization methods: attachment to solid support, or entrapment into gel particles. The first group is more applicable in practice, because of the lower mass transfer resistance. It comprises in binding the cells to solid surfaces by weak van-der-Waals bonds (flocculation, adsorption), ionic attraction, or by strong (chemical) bonds: covalent bonds or cross-linking. However, due to the completely different size and environmental parameters of the cells, the relative importance of these methods is considerably different. The criteria imposed for cell immobilization technique usually determine the nature of the application. Adsorption is the first example of cell immobilization. Adsorption is based on electrostatic interactions between the charged support and microbial cell. However, the actual charge on support surfaces is still unknown and this limits the proper choice for microbial attachment. Along with charge on the cell surface, the composition of cell wall carrier composition will also play an important role. Carrier properties will also greatly influence cell-support interaction. The trend is to combine the adsorption properties (strength of the bonds, highly developed adsorption surface) with the carrier price. Therefore there are many efforts to use relatively cheap carriers, like glasses or ceramic supports as well as varying proportions of alumina, silica, and oxides of magnesium, zirconium, etc. A variety of microbial cells were immobilized by adsorption on different supports [2] like kieselguhr, wood, glass ceramic, plastic materials, etc. Hattori&Furusaka [3, 4] reported the binding of Escherichia coli cells onto an ion exchange resin. There is some data about magnetic retention of cells onto a carrier [5].
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The mechanism involved in covalent binding is based on covalent bond formation between activated support and cell in the presence of a linking agent. For covalent linking, chemical modification of the surface is necessary [6]. The covalent bonding agent is selected depending on the properties of the cell membranes. There are different methods for cell covalent binding to a solid support depending on whether the microbes are Gram-positive or Gram-negative ones. Covalent binding and cross-linking offer better strength, than the adsorption or flocculation but there is an encountered toxicity in the reagents used to produce immobilization [7]. Microbial cells can be immobilized by cross-linking with bi- or multifunctional reagents such as glutaraldehyde [8] and toluene-di-isocyanate [9] after proper modification of the carrier surface. There are also other physical processes, such as flocculation [10] and pelletization [11]. They are based on the strong mutual adherence forces of some microbial cell cultures. There are results for bacteria immobilized on poly (acrylo-nitrile-acrylamide) [12], cellulose [13], etc. The first group, i.e. fixation of microbial cells onto solid support has found more practical applications. The main reason why is the lack of, or the low mass transfer resistance for substrates penetration or products removal from the space around the cells. The application of the first group of methods usually leads to formation of ‘biofilm” on the surface of the solid support. The second group of methods is based on physical retention (or entrapment) of cells in the structures of gels (ionotropic gelation, thermal gelation polymerization) or membrane retention (dialysis culture, ultrafilters) cf. [14]. The most frequently used gels are polyacrylamide [15-21], calcium alginate [22], carageenan [23-26], agar [27], and their modifications. Cell entrapment in gels, or retention by membranes has been and it is still extensively studied but it has found some medical applications mainly. A successful industrial application for L-aspartic acid production by cells immobilized in gel (poly-acrylamide) was reported in 1983 [28]. The most widespread method of this group is entrapment in calcium alginate. It was applied for production of antibiotics [29-31] and organic acids [32-34]. Since the production of acids leads to acidification of the broth, calcium alginate may dissolve and the biocatalyst may be destroyed. There are some indications, that in some cases cells entrapped in gel of κ-carageenan give higher product yields compared to the attached ones [35]. There are bioreactors used for wastewater treatment with bacterial cells entrapped into gel beads [26, 36, 37]. An excellent and comprehensive review of immobilized cells, methods and applications is made by Ramakrishna&Prakasham [38].
III. Biofilm Reactors The simple definition of a biofilm is “microorganisms attached to a surface.” Actually biofilms are a complex aggregations of microorganisms growing on a solid substrate. Biofilms are characterized by structural heterogeneity, genetic diversity, complex community interactions, and a self-produced extra-cellular matrix of polymeric substances [39]. In general, there are four stages to the development of a mature biofilm: initial attachment,
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irreversible attachment by the production of extracellular polymeric substances, early development, and maturation of biofilm architecture [40]. Formation of a biofilm begins with the attachment of free-floating microorganisms to a solid surface. The first cells adhere to the surface initially through van der Waals forces. If they are not immediately separated from the surface, they can attach themselves using cell adhesion molecules. Then they facilitate the arrival of other cells by providing more diverse adhesion sites and beginning to build the matrix that holds the biofilm together. Other species are able to attach themselves to the matrix or directly to earlier colonists. Then the biofilm grows by cell division and recruitment [41]. Biofilms are usually found on solid substrates submerged in or exposed to aqueous media. Biofilms consist of many species of bacteria living together within a matrix of excreted polymeric compounds (EPC). This matrix protects the cells and enables communication among them through chemical and physical signals. Some biofilms have porous structure with water channels that facilitate penetration of nutrients and signalling molecules. Biofilm is important survival mechanism for bacterial cells. The cells can avoid attack by host defenses and to resist toxic compounds at higher concentration than in free culture. For example, disinfection through chlorination of a biofilm is usually unsuccessful because the biocide only kills the bacteria in the outer layers of the biofilm. The bacteria within the biofilm remain safe, and later the biofilm can restore itself. Such effects have been demonstrated and studied for the resistance of cells in biofilm to antibiotics [42, 43]. Within a biofilm, a variety of microbial groups can contribute to the conversion of different organic and inorganic substrates. For example, when a wastewater contains a mixture of conventional and xenobiotic organic pollutants, biodegradation of the xenobiotics requires a population of slow-growing organisms – those capable of degrading the xenobiotics. The latter are easily washed out of a suspended-growth process, since all the biomass has the same growth rate, whereas the slow-growing bacteria can establish themselves deeper inside the biofilm, protected from loss, while the conventional pollutants are removed near the biofilm-fluid interface [44]. There are a plenty of research papers on biofilm applications, particularly in wastewater treatment. Some of they are related to nitrification/dentrification of wastewater [44-46] or ions retention [45-50]. The most extensive applications are the biodegradation of oil residues and derivarives [51, 52], xenobiotics (phenol on a first place [53, 54]) and other organics [55]. In anaerobic waste water treatment, these reactors have been used at full scale successfully throughout the world, mainly in food and beverage industries [56]. The industrial large-scale applications of biofilms are in batch reactors, continuous stirred tank reactors (CSTR), packed bed reactors (PBR), trickle-bed reactors (TBR), fluidized bed reactors (FBR), airlift reactors (ALR), upflow anaerobic sludge blanket (UASB) reactors. PBRs are packed with suitable support material followed by inoculation with the culture to form biofilm. Such reactors are subjected to clogging and blockage due to excessive cell growth without cell detachment because of the stagnant particle layer. Very high reaction rates are attained compared to batch reactors [57].
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TBRs are fed by liquid phase at the top of the reactor thus obtaining product at the bottom. However, in such reactors some of the biofilms may not get sufficient feed because of the non-uniform fluids distribution, thus affecting reactor efficiency/productivity adversely. This also may affect the efficiency of the reactor. The most widespread biofilm reactor is the biofilter [58, 59]. Biofilters are effective and mostly used for the treatment of wastewater and air, polluted by noxious or hazard chemicals to convert them into harmless end products. Such pollutants usually are hydrogen sulfide, mercaptanes, volatile organic compounds, and odors from natural and household sources. Biofilters are assembled as trickle-bed reactors with up-flow gas feed and down-flow water feed with biomass attached to particles in a packed bed. FBRs have played a successful role in the degradation of toxic phenolic chemicals [60] and butanol production [61, 62]. A major advantage in these reactors is that they can be operated for much longer periods than PBR or CSTRs. They have much lower pressure drop than PBR and they do not block due to excessive growth. Fluidized bed reactors (FBR) are operated with up-ward water flow that suspends small carrier particles in the water phase. The biomass is subjected to lower shear stresses compared to CSTR. In airlift reactors (ALR), the carrier particles are suspended in the circulating water flow that is caused by the injection of air. With suspended support media, the moving bed biofilm reactor does not have to be backwashed as biofilm detachment is caused by particle-particle collisions within the system. Airlift reactors (ALR) contain two concentric tubes, a riser (an inner tube) and a downcomer (an outer tube). In these reactors, mixing is achieved by circulating air introduced at the bottom of the inner tube. Such reactors where air is replaced by an anaerobic gas are called gas/lift reactors. They are used for different purposes, for biodegradation of organics as well [63]. The most commonly used bioreactors that fall in the category of the biofilm reactors are the upflow anaerobic sludge blanket (UASB) reactors. The UASB reactor was developed by Lettinga et al. [64]. They are mainly used for anaerobic treatment of domestic and industrial wastewater. The liquid or slurry is introduced in these reactors in upward direction through a “blanket” of immobilized cells or floccules. Reactor effluent and the produced gas are removed from the top of the reactor. UASB has been used for biodegradation of some xenobiotics as well [65, 66]. A separate class of biofilm reactors is the rotating biological contactor (RBC), consisting of horizontal discs with immobilized biomass on them, mounted on a horizontal shaft and partially dipped into a pool with the treated liquid. The advantages for this bioreactor type are the simplicity of maintenance and operation, the low power consumption, the ability to withstand shock or toxic loads and the sludge settling properties. The rotating biological contactor process was first used in Germany in the 1920's. It is extensively used for large scale wastewater treatment [67, 68]. Typical applications of RBCs are treatment of: municipal wastewater, food & beverage wastewater, landfill leachate, refinery & petrochemical wastewater, pulp & paper wastewater, septage and industrial wastewaters. They were commercialized for many decades for wastewater treatment in wine distilleries [69], mining industry and ore dressing [70, 71], municipal wastewater [72], etc.
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The kinetic study of industrial microbial processes in biofilms includes the evaluation the contribution of the cells attached to the solid support and the contribution of the free cells detached from the particles and growing as free ones in the liquid. The cell detachment process is very important for the operational stability and multiple use of immobilized biocatalysts. Therefore it is interesting to know the share of cells detached from the carrier and the detachment rate compared to the rate of microbial growth within the biofilm. There are papers with evidences that steady-state in biofilm biomass concentration is never reached because of the periodical way of existence and the different periods of growth - initial attachment, irreversible attachment, early development, and maturation followed and accompanied by cell detachment and the biofilms are un-reproducible [73]. Cells are separated periodically from the carrier and are washed out by the stream. Therefore an oscillation regime in biomass detachment takes place [74].
IV. Modeling of Biofilm Performance This matter has been systematically and comprehensively developed by a Task Group at the International Water Association [75]. The mathematical modeling may have different goals. In the case of biofilm the chosen model should match the following goals as much as possible. 1. First of all it is the understanding of fundamental mechanisms - how a biofilm forms and performs. This is accomplished through comparison of mathematical models results with experimental data and test for model adequacy. 2. To integrate different processes occurring in a different spatial and temporal scale: transport processes, metabolism, chemical, mechanical processes, etc. 3. Modeling gives researchers and practitioners the opportunity to select the most promising designs for experimental testing and further development for practical applications.. 4. To improve the performance of a process, either new, or running ones using the predictions of adequate models. The systematic way that the parts of a model are built up so that they represent the desired feature: •
• • •
divide the space that includes the biofilm into distinct zones (or compartments), such as the carrier onto which the biofilm accumulates, the biofilm itself, the concentration boundary layer around the biofilm and the overlying liquid; outline characteristic dissolved components, such as substrates, and particulate components, i.e. bacteria; consider the transport and transfer processes to supply substrate, nutrients and oxygen to the cells in the biofilm; define the rate of reactions that have quantifiable parameters;
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•
compose mass balance equations including the rate terms for all the processes that affect a component in a particular spatial compartment; solve the mass balance equations by a numerical or analytical technique, yielding the model output; evaluate model parameters by comparison with experimental data, i.e. through identification procedure by minimizing objective function, i.e. sum of least squares between model and experimental values; distinguish among different models in ways that are relevant for selecting a model. Test for adequacy.
There are different classifications of mathematical models, particularly for biological systems. Depending on the mathematical method for solution of the equations they are analytical and numerical. From spatial point of view, they could be one-, two-, or threedimensional. Multidimensional numerical models: The biofilm is considered as two- or threedimensional structure. All concentrations (of substrate, products), a well as the biofilm structure and properties can vary in multi-dimensional space and they are time-dependent. Numerical solutions require more computing power than for one-dimensional models, but in general, they are now feasible. One-dimensional numerical models: All quantities are averaged in the plane parallel to the solid support, but gradients perpendicular to this support can be calculated for all components. When a spherical symmetry exists, the one-dimensional models give sufficient spatial presentation. Analytical models include simplifying assumptions so that the flux of dissolved substrates into the biofilm can be calculated easily. For this end the model equations should be linearized. In his case numerical treatment is needed for tabulation of special functions and to avoid tedious manual calculations. They are applicable for a single species and substrate, but these models are of limited value. Moreover, parameter evaluation always requires minimization of objective functions including sum of differences of experimental and model values. In these procedures the use of numerical methods is inevitable. That is why the analytical models are of little use. An important problem arises when many model parameters have to be evaluated, from experimental data by minimizing of the objective function. Sometimes best fit is observed at parameter values with no physical meaning. In other cases the objective function is not sufficiently sensitive to some parameters. That is why the more parameters are previously known, the more reliable the modeling results are. Another tricky issue is the appropriate selection of initial approximations for the parameters when starting the minimization procedure. The main uncertainty in parameter selection comes from the fact that the essential model parameters can be different for the liquid phase and for the biofilm or the polymer matrices. Things become more complicated when more microorganisms, mostly unknown, are attached and involved in the overall microbial process as consortia [76,77]. This is the usual case, particular in waste water treatment. The detailed description of all processes and the
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interactions between the cells in the biofilm is too complicated to be formulated accurately, if known at all. Systemized approach for these issues is given in [75]. Mechanistical modeling of biofilms is also applied [78, 79]. These mathematical models represented the biofilm as a well defined shape, i.e. simple “slab” or “plate” in which substrate gradients are in one dimension, perpendicular to the surface onto which the biofilm is attached. As a rule, the biofilm is considered as homogeneous structure in this approach. These models are based on experimental measurements of the overall substrate-removal rate and the total biofilm accumulation. Nowadays the advanced experimental techniques enable deeper insight into the biofilm structures and behavior. Microsensors can be used to measure concentrations directly within the biofilm [80-82] (e.g., oxygen, ammonia, nitrate, pH). Recent advances in molecular biology and in situ hybridization techniques have resulted in the development of gene probe, fluorimetry and different microscopy techniques that enabled the detailed analysis of biofilms structure and behavior of the microogranisms within it [81-86]. The newly accumulated information on biofilms led to more comprehensive knowledge and therefore to the development of new, more detailed mathematical models. The visualization of heterogeneous structures in biofilms has prompted the development of a new generation of mathematical models in which the three-dimensional structure of the biofilm is simulated. A multi-species biofilm analytical model has been developed by Wanner&Gujer [87]. It takes into account the mass transfer effects in the biofilm, the external mass transfer and predicts the variation in film thickness as well as the spatial distribution of microbial species and substrates. The model implies cell detachment from the biofilm due to shear forces or sloughing. Picioreanu et al. [88] proposed a two-dimensional model of biofilm detachment by shear stress as well as by cell deplacement caused by microbial growth. Van Loosdrecht et al. [89] analyzed the mathematical models on biofilm growth, biofilm properties and microbial detachment from the carrier. They have shown that the morphology and the strength properties of biofilms strongly depend on the shear stresses the biofilm particles are exposed to, on the microbial growth and on the spatial substrate distribution and concentration gradients. Under certain conditions the biofilms might be smooth, uniform and stable (at higher shear stresses, low specific rate of growth and uniform substrate concentration within the film) or heterogeneous, unstable and patchy-like (at high specific growth rate and large concentration gradients). Recently a steady model for the evaluation of external liquid film diffusion and internal pore diffusion effects in an immobilized biofilm system under continuous mode of operation was developed [90]. The model takes into account, substrate diffusion through external liquid film and biofilm. Average rate of substrate consumption in the biofilm was considered. The overall efficiency of the biofilm was mathematically represented by considering the combined effects of substrate penetration and substrate utilization in the biofilm. The model is able to effectively predict both internal and external mass transfer effects in an immobilized biofilm system. The model was illustrated using a case study of pyridine biodegradation in a rotating biological contactor immobilized with pyridine degrading microbial film. A model for fluid flow, and contaminant and nutrient transport, coupled with biofilm growth was proposed [91]. The model is takes into account the effect of the flow on the biofilm growth and vice versa, so that all the effects are strongly coupled.
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V. Processes on Granulated Activated Carbon (GAC) A special separate case of biofilm reactor is the one operating with biomass adsorbed on granulated activated carbon (GAC) [92, 93]. In this case two important properties of GAC are used: first the microbial cells are easily attached to the carbon surface; on the other hand the toxic compounds to be treated are adsorbed by the activated carbon. The toxic substrate concentration in the bulk is reduced to values affordable for the cells and the cells are protected from the shock loads of organic pollutants [94-96]. Therefore the cells are not subjected to the adverse action of the toxic substrate or inhibitor at high concentrations. After biodegradation when the dissolved toxic substrate is degraded, new portions of it are released by the carbon because of the adsorption/desorption equilibrium. That is why the degradation rate for immobilized cells is usually much higher than that for free ones. However, there could be a competition for the adsorption sites between the substrate molecules and the microbial cells, resulting in apparently lower adsorption capacity [97-99]. Herzberg et al. [100] claimed that the biofilm coverage has a patchy-like nature enabling the simultaneous adsorption of solutes and cell attachment. They confirmed this explanation with own experimental data for atrazine biodegradation by Pseudomonas cells in biofilm on GAC. After regeneration, the GAC is no longer considered a hazardous waste and may be reused. However, regeneration reduces the adsorptive capacity of GAC and the used material eventually must be disposed of and replaced. GAC consumption is dependent upon flow rates and contaminant concentrations. Aqueous phase GAC can be subject to clogging from carbonate precipitation or biofouling, therefore, pretreatment of the influent water stream may be required. Generally, GAC is cost effective for low flow and low concentration applications [101, 102].
Concentrations, mg/l
6000
4000
2000 inlet concentration exit concentration 0 0
20
40
60
80
100
120 140 Time, h
Figure 1. Biodegradation of aniline in continuous fixed bed reactor with biofilm of Pseudomonas putida, fixed on granulated activated carbon (own not published data). Dilution rate – 0.17 h-1.
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1800
Concentrations, mg/l
1500 1200 900 600
inlet concentration outlet concentration
300 0 0
50
100
150
200
Time, h
250
Figure 2. Biodegradation of nitrobenzene in continuous fixed bed reactor with biofilm of Pseudomonas putida, fixed on granulated activated carbon (own not published data). Dilution rate – 0.2 h-1.
There are different applications of GAC for degradation of chlorinated compounds [103107], of phenol and its derivatives [98, 99,108-112], aromatic hydrocarbons [113], etc. There are different attempts to describe the removal of organic pollutants by GAC by adsorption only [114] or by simultaneous adsorption and biodegradation [115-118]. Pilot-plant data for biodegradation of aniline and nitrobenzene by free and cells of Pseudomonas putida adsorbed on GAC is shown in Figures 1, 2 (own not published data). In both cases the biodegradation rate was higher than 99%, whereas the process could not start in free culture. The granulated activated carbon method has been commercially applied for wastewater treatment in the 1980s as “Biocarbon” process. Papers on mathematical modeling of xenobiotics biodegradation by cells on granulated activated carbon are known in the recent literature [115-118]. The unstructured model in [115] is based on the model concept of one biodegradable and one non-biodegradable component of complex mixtures and the competition of physical adsorption and biodegradation. It provides a good screening tool for initial assessments of process feasibility, preliminary economic analyses and planning of detailed experimental studies. A mathematical model of the biodegradation of xenobiotics by microbial cells attached to particles of granulated activated carbon was developed in [118]. It takes into account the substrate adsorption on the activated carbon. The model allowed the quantitative evaluation of different characteristics of the biofilm behavior: retarded microbial growth, increased concentration of immobilized cells compared to suspended cultures, potential cell detachment from the solid support and consequent independent growth of free cells. The applicability of the model was demonstrated for our own experimental data for 1,2-dichloroethane (DCA) biodegradation by Klebsiella oxytoca VA 8391 cells attached to granulated activated carbon. Two types of reactors: recirculated batch and continuous flow bioreactor were studied. It was shown that in both investigated cases - recirculated batch reactor and continuous flow reactor,
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the major contribution to DCA biodegradation was provided by immobilized cells, whereas the cell detachment rate was negligible. The immobilized cells were found to tolerate much higher substrate concentration and much higher dilution rates in continuous culture than the free cells.
Vi. Processes in Entrapped Cells There are observations for enhanced stability and resistance of entrapped cells toward toxic substrates. Kobayashi&Nakamura [119] reported that the immobilized cell culture had higher ethanol productivity than the free cells and produced ethanol continuously with a dilution rate of 0.5 h−1, which is about 10 times larger than the washout rate in a free cell culture. Chen et al. [120,121] have found that the strain Klebsiella oxytoca was able to biodegrade cyanide to non-toxic end products at high level of KCN concentration and wide ranges of pH when entrapped cells have be applied, whereas in free suspension systems the cell viability was highly affected by initial KCN concentration and pH. The main problem however that impede the larger application of entrapped cells in practice is the considerable mass transfer resistances for substrate supply and product removal because of the slow molecular diffusion within the gel membranes and pores. The problem of mass transfer resistance in heterogeneous catalysis science is not subject to biocatalysis only. It has been extensively studied since the 1930s, for evaluation the decrease of catalytic activity due to the low diffusion rate within the pores of solid catalysts. In general, the chemical reaction accompanied by diffusion in porous (but considered as pseudo-homogenous) media can be described by the following set of partial differential equation (1) with the appropriate initial and boundary conditions (2):
∂C S = DS ∇ 2 C S − VS ∂t ∂C P = DP ∇ 2 C P + VS , ∂t
(1)
t = 0, C S = C S0 , C P = C P0 ; ∂C S ∂C P = = 0; ∂r ∂r r = R, ∂C S − DS = k L (C S − C S ,∞ ), ∂r ∂C P − DP = k L (C P − C P ,∞ ). ∂r r = 0,
(2)
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With D the diffusivities of the species are denoted. Indexes “S” and “P” denote the substrate and the product respectively and the particle radius is denoted with “R”. With “∞” the quantities related to the bulk phase far enough from the particles are denoted. For convenience, spherical particle symmetry is chosen. After normalizing the equations (1, 2) the following dimensionless parameters are deduced:
VSmax R 2 Φ = 0 , C S DS 2 S
k R Bi = L . DS
(3)
The first one known as Thiele modulus, represents the ratio between the time for diffusion ~R2/DS of substrate and product molecules and the time for chemical reaction ~CS0/VSmax and among the pores. The second one is the Biot number representing the ratio of the rates of external mass transfer and the diffusion rate within the particles. At Bi>>1 the external mass transfer resistance is negligible. This condition is usually satisfied at Bi>10. The reaction efficiency ξ, i.e. the ratio of the reaction rate in porous medium V to the intrinsic one (V0) in homogeneous medium in a well mixed state, depends on the Thiele modulus [122]:
ξ=
V = f (Φ S2 ) V0
(4)
The higher Φs2, the lower the efficiency because of the stronger diffusion limitations. For the case of enzyme reactions following the Michaelis-Menten kinetic equation the Thiele modulus Φs2 is usually written in the form [123,124]:
Φ 2S =
Vmax R 2 . K m' Deff
(5)
In the case of immobilized microbial cells however, it is not so easy to define the Thiele modulus. It is because of the cell growth, the viable cell concentration, depending on time and the complicated cell metabolism under these conditions: impeded microbial growth, mass transfer limitations and odd cell distribution within the pores. That is why, the Thiele modulus should be carefully defined for each separate case. For the case of immobilized cells, able to grow within the gel particles Eq. (1) takes the form:
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∂C S = DS ∇ 2 C S − VS ∂t ∂C P = DP ∇ 2 C P + VS , ∂t VS = f (C S , C P , X im )
(6)
dX im / dt = μ im X im − γX im With γ the specific rate of cell death is denoted. If entrapped in gel, the immobilized cells can grow within the particles. The cells situated at the particle periphery are in a favorable situation because they are close to the adjacent liquid phase and they are well supplied with substrate, nutrients and oxygen. The cells far below the particle surface suffer of limited supply and therefore they growth will be impeded. Additionally accumulation of products of the reactions, due to mass transfer limited removal being possibly inhibitors will worsen the situation [125-127]. Mass transfer limitations may cause reduced oxygen supply and uptake rate and non-sufficient substrate and nutrient supply for the cells with the resulting altered cell metabolism [128, 129]. Another reason for impeded growth is the spatial limitations within the pores [130-136]. Because of all these effects of mass transfer limitations it is generally assumed that the immobilized cells are concentrated in a thin layer close to the particle/liquid interface even when the cells are initially uniformly distributed after immobilization [127-141]. This statement was confirmed by experimental data [133, 134, 138-140]. Other complications are the different diffusivities in gel pores compared with those in solutions [141-158] and the influence of the immobilized cells on the diffusion coefficients of the solutes [141,149, 150]. There are different papers, dedicated to these problems with mathematical modeling the processes of impeded microbial growth coupled with substrate and product diffusion in the particle pores [151, 152, 149-158]. In general all of these papers suggest an odd cell distribution. Some of them are qualitatively supported by experimental results for lactic acid production in continuous culture [141]. However, there were observations in the latter paper, that transitions to a new pseudo-steady state, after changing dilution rate were much slower in the experiment than those predicted by the model. Therefore these authors stated that “an accurate description of immobilized cell activities cannot be done with kinetic models for free cells, … due to a change in the physiology of immobilized cells” [141]. This statement was based on other studies claiming that such discrepancies with the behavior of free cells may be due to changes in cell growth and physiology [159], like changes in the redox states, enzymatic pool and intra-cellular pH [160], shift in the metabolic pathway from homofermentative to hetero-fermentative and increased tolerance to environmental stresses [161, 162]. Significant pH-reduction within the particles was observed because of lactic acid accumulation at lactic acid fermentation [158]. There are also other data about alterations in cell physiology after cell immobilization [163, 164]. Actually, the cell profile along the particle radius is much steeper in practice than the models predictions and the cells are concentrated within a thin layer close to the particle surface [133, 134, 139, 140].
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This facts were taken into account in mathematical modeling the processes by immobilized cells entrapped in polymer matrices or attached onto polymer carrier [154, 165168], where different values for maximum specific growth rate and yield coefficients for immobilized cells have been assumed, but at the same Monod structure for the kinetic equation for specific growth rate The mathematical models based on Eqs. (5) were used to evaluate these parameters for immobilized cells from experimental data for gluconic acid [165], lactic acid [166, 167] and CGTase production[168]. The estimated values for μmax from own experimental data by the models are shown in Table 1. Review of these data for the maximum specific growth rate shows one clear trend. The Lactobacillus casei cells attached to polyurethane with large pores can grow like the free ones with an equal μmax. The results for Gluconobacter oxydans: show that the immobilized cells have similar, although somewhat lower specific growth rate than the free ones. In the case of CGTase production – the maximum specific growth rate for the immobilized cells attached onto poly-sulfonate support is of the same order of magnitude as for free ones but it is twice as much lower. In these three cases there is almost no spatial hindering for the cells and their growth. The case of lactic acid fermentation by cells L. rhamnosis entrapped in polyacrylamide gel with narrow pores shows that their growth is strongly impeded – the maximum specific growth rate for immobilized cells is 28 times lower than for the free ones. It is generally considered that accumulation of product within the gel membranes leads to retardation of microbial growth and production provided the product is an inhibitor. Therefore for such processes cell entrapment should be avoided as immobilization method. Another possible case is the different yield coefficients for immobilized cells. This case was considered in [168] for CGTase production by Bacillus circulans cells immobilized onto poly-sulfonate membranes. It was demonstrated by mathematical modeling and the experimental data that in the considered case the product yield coefficient is much higher than for free cells. Table 1. Comparison of maximum specific growth rate of immobilized cells estimated by mathematical models from experimental data with the ones for free cells Process
μmax, -1
Gluconic acid production, Gluconobacter oxydans, Caalginate, substrate and product inhibition assumed [165] Lactic acid production, Lactobacillus rhamnosis, PAA, substrate and product inhibition assumed [166] Lactic acid production, Lactobacillus casei, polyurethane foam, product inhibition assumed [167] CGTAse production, Bacillus circulans, cells attached onto poly-sulfonate membranes, substrate inhibition assumed [168]
μmax, h-1
h free cells 0.43
immobilized cells 0.37
0.56
0.02
0.33
0.33
2.0
1.0
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However, when the bio-reaction is an auto-catalytic one, cell entrapment could be favorable because of accumulation of product in the gel beads caused by the low removal rate. This effect was demonstrated for the case of D-sorbitol to L-sorbose biotransformation by cells of Gluconobacter oxydans entrapped in poly-acrylamide gel [19]. In this case the mathematical model (1, 5) with the related kinetic equation was written in the following dimensionless form with the initial and boundary conditions (2): ∂C S = ∇ 2 C S − Φ S2 C S C 2P ∂T ∂C P = Rd∇ 2 C P + Φ S2 C S C 2P ∂T . Φ S2 = kR 2 /D S ,
(7)
Rd = D P / D S . η=
CP C + C 0P 0 S
The degree of conversion η is defined as a ratio of the product concentration to the sum of initial concentrations of substrate and product. Some of the results of the solution of Eqs.( 7) are shown in Figure 3. The effect of the diffusivity ratio Rd (product to substrate diffusivities) on the degree of conversion is very important, particularly when the product is inhibitor, or catalyst, as it is the discussed case. The lower the diffusivity ratio, the stronger the autocatalytic effect. The degree of conversion will higher compared to free culture. Comparison of experimental data with the model predictions is shown in Figure 4.
Figure 3. Model (7) results for variation in conversion η with time at different diffusivity ratios for autocatalytic reaction [19]. Line 1 – Rd = 1.5; Line 2 – Rd = 1; line 3 – Rd= 0.5. Bi = 1000; Φs2 = 1.
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Figure 4.Comparison of experimental results (●) with the model (line) for autocatalytic biotransformation (D-sorbitol to L-sorbose). Symbols (○) – reference experiment with free cells [19].
VII. Cell Growth and Detachment from Gel Particles As the cells can grow in the particles or in the biofilm, their concentration can become so high that they could disrupt the particles or simply to leak into the liquid medium [151, 154, 165-169]. The detached cells can grow and produce in a free culture contributing to the overall fermentation process. The operational stability of the biocatalyst and its multiple use depends on the balance between the rates of cell growth and cell detachment. The cell leakage may result in biocatalyst exhaustion provided the leakage rate is faster than microbial growth. There are efforts for quantitative evaluation of this effect introducing a cell leakage factor, accounting for the rate of cell detachment from the particles [165-168]. The balance for the cell in free culture with cell detachment from the particles is given by:
dX = μX + β a[μ im X im ]r = R , dt t = 0, X = X 0
(7)
Here a is the specific interfacial area of the particles, β is the cell detachment factor and the indexes denote the quantities for the immobilized cells. The microbial growth within the particles was described by the equation, following the Monod type kinetics with product and/or substrate inhibition:
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Table 2. Estimated values of the dimensionless leakage factor β = β / R , [h-1] from experimental data for consecutive runs [166]
Gel A Gel B Gel C Gel D
Run 1 1.10-8 2.10-9 2.10-8 2.10-10
Run 2 0.011 0.005 5.10-4 0.008
Run 3 0.065 0.01 0.03 0.005
Run 4 3.2.10-4 1.1.10-4 5.10-4 0.001
o X im = X im . exp(μ im t ) ,
(8)
assuming uniform cell distribution as an initial condition. The cell detachment factor β could be considered as a thickness of the layer emitting cells from the particle surface into the broth. If β = 0, no cell detachment takes place, no free cells grow in the broth and there are only the immobilized cells which contribute the fermentation process. However, if β = β / R is close to unity i.e. β ~ R, the culture will behave like a free one and the particles will be exhausted, sooner or later. Experimental studies and modeling on this phenomenon has been made in [154, 165168]. The values of the estimated leakage factor for lactic acid fermentation by cells of Lactobacillus rhamnosis entrapped in poly-acrylamide gel [166] are given in the Table 2. It is demonstrated that this leakage factor and therefore the biocatalyst operational stability depends on the way of gel preparation and on the consecutive run. Gels A-D have been prepared under different conditions with different void fractions and strength. The pores size decreased from Gel A to Gel D. Reviewof these data shows that in all of the first runs the leakage was negligible. This is due to the low initial cell concentration and the very thin, almost negligible layer at the particle/liquid interface where the cells are. During the first runs the fermentation is accomplished only by the immobilized cells for each gel. The rapid increase of β for the next runs shows, that the cell concentration within the particles becomes sufficiently high to facilitate cell leakage from larger layers close to the particles surface. The further decrease of
β shows that the particles are already exhausted. These processes are better expressed for gels with wider pores and relatively large void spaces like Gels A and B. The best operational stability is demonstrated by Gel D, where the dimensionless leakage factor β remains low and stable for the consecutive runs. Similar analysis has been made for the productivity of cells attached onto poly-sulfonate membranes for CGTase production [168]. Mathematical modeling in this case has shown that the kinetic curves are very sensitive to the cell leakage. That is why the contribution of the immobilized cells and the free ones developed in the broth after detachment can be easily distinguished. This mathematical modeling makes it possible to estimate the cell detachment rate and to separate the contribution of free and immobilized cells to the overall fermentation process.
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VIII. Conclusion The analysis of the data in literature show that although the great scientific interest to the performance of immobilized cells their industrial applications in chemical technology and biotechnology are restricted mainly to some specific processes, like waste water treatment. Broader applications have the cells, attached in form of biofilm on solid supports, granulated activated carbon, in particular. The main problems for the practical application of microbial cells, immobilized in biofilm or entrapped in gels are the mass transfer resistance resulting in substrate concentration gradients and limited supply and the spatial hindrances for cell growth and supply with substrate and oxygen for the cells inside the particles. That is why the cells immobilized in gels are concentrated in thin layer close to the particle surface. On the other hand, immobilized cells show enhanced operational stability and resistance toward toxic substrates and substrate-inhibitors at higher concentrations. Much higher dilution rates in continuous culture can be attained in comparison to free cells. The operational stability of biofilms is strongly dependent on the shear stress in the fluid flow, on the substrate supply and on the microbial growth rate. This operational stability is strongly influenced by the mechanical properties of the biofilms and the gels, as well as by the cell detachment and leakage from the particles. The comparison of experimental data with unstructured mathematical models shows that usually cell behavior and the kinetic constants for cell growth and production are different from those for cells in free culture. There are evidences that immobilization in gels and biofilms may lead to physiological changes in cell function. The main research area on immobilized cell operations will be to study experimentally the cell behavior under the conditions of accumulation in the limited space environment and the resulting altered physiology and metabolism. Then more realistic and reliable mathematical models can be developed to quantify the processes with immobilized cells and to apply them in practice.
References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10] [11] [12]
Chibata, I.; Tosa, T.; Sato T. Appl Microbiol 1974, 27, 878–885. Klein, J.; Ziehr, H. J Bacteriol. 1990, 16, 1–16. Hattori, T., Furusaka, C. J Biochem 1960, 48, 831–837. Hattori, T., Furusaka, C. J Biochem 2001, 50 312–315. Safarik, I., Safarikova, M., China Particuology, 2007, 5, 19-25. Novarro, J. M., Durand, G., Eur J Appl Microbiol 2001, 4, 243–254. Kolot, F. B. Process Biochem 1981, 16, 2–9. Chibata, I., Tosa, T. and Sato, T. 1974, Japanese Patent No. 7,425,189. Lartigue, D. J. and Weetall, H. 1976, U.S. Patent No. 3,939,041. Lee, G. K., Long, M. E. 1974, U.S. Patent. No. 3,821,086. Frick, Ch., Schügerl, K. Appl Microbiol Biotechnol 1986, 25, 186-193. Lalov, I. G., Krysteva, M. A., Phezoulat, J.-L. Bioresource Technol 2001 79 83-85.
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[13] Krysteva, M. A., Grigorova, P. M. Enzyme Microb Technol 1987, 9, 538-541. [14] Chibata, I. In Immobilized Microbial Cells; Venkatsubramanian, K., Ed.; ACS Symposium Series 106; American Chemical Society: Washington DC, 1979; pp 187202. [15] Mosbach, K., Mosbach, R. Acta Chem Scand 1966, 20, 2807–2810. [16] Schnarr G.W., Szarek W.A., Jones J.K.N. Appl Environ. Microbiol 1977 33 732-734. [17] Freeman, A., Aharonowitz, Y. Biotechnol Bioeng 1981, 23, 2747–2759. [18] Mittal, Y., Mishra, I. M., Varshney, B. S. Biotechnol Tech 1993, 15, 41–46. [19] V. Beschkov, V., Kosseva, M. Chem Eng J 1990, 45, B5-B11. [20] Stefanova, S., Koseva, M., Tepavicharova, I., Beschkov, V. Biotechnol Lett 1987, 9 (1987) 475-477. [21] Petrov, K.K., Petrova, P.M., Beschkov, V.N. Bulg Chem Commun 2005 37, 173-177. [22] Kierstan, M., Bucke, C. Biotechnol Bioeng 1977, 29, 387–397. [23] Chibata, I., Tosa, T. Ann Rev Biophys Bioeng 1981, 10, 197-216. [24] Chibata, I., Tosa, T., Sato, T. Appl Biochem Biotechnol 1986, 13, 231-240. [25] Nishida, Y., Sato, T., Tosa, T., Chibata, I. Enzyme Microb Technol 1979, 1, 95−99. [26] Hunik, J. H., Bos, C.G., van den Hoogen, P., de Gooijer, C. D., Tramper, J. Biotechnol Bioeng 1994, 43, 1153–1163. [27] Vassileva, A., Beschkov, V., Ivanova, V. Tonkova, A. Proc Biochem 2004, 40, 32903295. [28] Chibata, I. In Basic Biology of New Developments in Biotechnology; Hollaender, A., Laskin, I., Rogers, P.; Eds.; Plenum Press, New York, NY, 1983; pp. 465−496. [29] Dalili, M. and Chau, P. C. Biotechnol Lett 1988, 10, 331–336. [30] Mahmoud, W., Rehm, H. J. Appl Microbiol Biotechnol 1987, 26, 333–337. [31] Mahmoud, W., Rehm, H. J. Appl Microbiol Biotechnol 1987, 26, 338–341. [32] Kurosawa, H., Ishikawa, H., Tanaka, H. Biotechnol Bioeng 1988, 31, 183–187. [33] Cachon, R., Molin, P., Divies, C. Biotechnol Bioeng 1995, 47, 567–574. [34] Saucedo, J. E. N., Barbotin, J. N., Thomas, D., Appl Microbiol Biotechnol 1989, 30, 226–233. [35] Yasouri, F.N., Foster, H.A., Biotechnol Techn 1991, 5, 149-152. [36] Cho, Y.-H., Knorr, D. Food Biotechnol 1993,7,115–126. [37] Nilsson I, Ohlson S. Columnar denitrification of water by immobilized Pseudomonas denitrificans cells. Eur J Appl Microbiol Biotechnol 1982, 14, 86–90. [38] Ramakrishna, S.V., Prakasham, R.S. Current Science 1999, 77, 87-100. [39] Ryder, C., Byrd, M., Wozniak, D.J. Curr Opin Microbiol 2007, 10, 644-648. [40] Stoodley, P., Sauer, K., Davies, D.G., Costerton, J.W. Ann Rev Microbiol 2002, 56, 187-209. [41] Derlon, N., Massé, A., Escudié, R., Bernet, N., Paul, E., Water Research 2008 (available on-line, doi:10.1016/j.watres.2007.11.016). [42] Stewart, P.S., Antimicrobial Agents Chemotherapy 1994, 38, 1052-1058. [43] Dibdin, G.H., Assinder, S.J., Nichols, W.W., Lambert, P.A., Journal of Antimicrobial Chemotherapy 38 (1996) 757-769. [44] Rittmann, B.E., Schwarz, A.O., Sáez, P.B. In Biofilms II; Bryers, J.; Ed.; John Wiley & Sons, New York, NY, 2000; pp. 207-234.
Immobilized Microbial Cells - Applications and Mass Transfer Phenomena
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[45] Liu, H.J., Yang, F.L., Zhang, H.M., Hu, S.W., Sun, C. Huan Jing Ke Xue 2007, 28, 2123-2128. [46] Antileo, C., Roeckel, M., Lindemann, J., Wiesmann, U. Water Environ Res 2007, 79, 1006-1014. [47] Cloirec, P. le, Andrès, Y., Faur-Brasquet, C., Gérente, C. Rev Environ. Sci. Bio/Technol. 2003, 2, 177-192 [48] Lahav, O., Green,M. Water SA 2000, 26, 51-58. [49] Nerenberg R., Kawagoshi Y., Rittmann B.E. Water Research 2008, 42, 1151-1159. [50] Quintelas, C., Fernandes, B., Castro, J., Figueiredo, H., Tavares, T. Chem Eng J 2008, 136, 195-203. [51] Al-Awadhi, H., Al-Hasan, R. H., Sorkhoh, N. A., Salamah, S., Radwan, S.S. Int Biodeter Biodegrad, 2003, 51, 181-185. [52] Farhadian, M., Duchez, D., Vachelard, C., Larroche, C., Water Res. 2008 (available on-line, doi:10.1016/j.watres.2007.10.021) [53] Viggiani, A., Olivieri, G., Siani, L., Di Donato, A., Marzocchella, A., Salatino, P., Barbieri, P., Galli, E. J Biotechnol 2006, 123, 464-477. [54] Daraktchiev, R., Kolev, N., Beschkov, V., Aleksandrova, T. Bioprocess Eng 1996, 16, 5-7. [55] Woolard, C.R., Irvine, R.L. Water Environ Res 1994, 66, 230-235. [56] Nicolella, C., van Loosdrecht, M.C.M., Heijnen, S.J. Trends in Biotechnology, 2000, 18, 312–320. [57] Qureshi, N., Schripsema, J., Lienhardt, J., Blaschek, H.P., World J Microbiol Biotechnol 2000, 16, 377-382. [58] VDI Richtlinien 3477. (1991) Biological waste gas/waste air purification, Biofilters. In VDI-Handbuch reinhaltung der Luft, Band 6, Verein Deutscher Ingenieure, Dusseldorf, Germany. [59] Pandey, R.A, Padoley, K.V., Mukherji, S.S., Mudliar, S.N., Vaidya, A.N., Rajvaidya A.S., Subbarao T.V. Bioresource Technol. 2000, 98, 2258-2267. [60] Tsuno, H., Kawamura, M., Somiya, I. Water Sci Technol 1996, 34, 335-344. [61] Qureshi, N., Maddox, I.S. Bioprocess Eng 1998, 3, 69-72. [62] Qureshi, N., Maddox, I.S. Bioprocess Eng 1991, 6, 63-69. [63] Gikas,P., Livingston, A.G. J Chem Technol Biotechnol 2006, 81, 858 –865. [64] Lettinga, G., van Nelsen, A.F.M., Hobma, S.W., de Zeeuw, W., Klapwijk, A. Biotechnol Bioeng 1980, 22, 699-734. [65] Sponza, D.T., Enzyme Microb Technol 2001, 29, 417-427. [66] Veeresh, G.S., Kumar, P., Mehrotra, I. Water Research 2005, 39, 154-170. [67] Submerged Aerated Filters (SAF); KEE Process Limited & KEE Services Limited,
[email protected]. http://www.keeprocess.com [68] Rotating Biological Contactors; GlobalSpec.com. http://process-equipment.globalspec. com/SpecSearch/Suppliers?Comp=3161 [69] Malandra, L., Lovis, K., Coetzee, G., Wolfaardt, G., Viljoen-Bloom, M. Evaluation of a rotating biological contactor for biological treatment of winery effluent, Wynboer. A Technical Guide for Wine Producers, http://www.wynboer.co.za/recentarticles/0204 rotate.php3
302
Venko Beschkov
[70] http://nrcan-rncan.gc.ca/mms/canmet-mtb/mmsl-lmsm/enviro/effluents/images/gallery/ pages/rbc_jpg.htm [71] Nikolov, L., Karamanev, D., Dakov, L., Popova, V. (2001) Environmental Progress 2001, 20, 247-250. [72] Hiras, D.N., Manariotis, I. D., Grigoropoulos, S.G. Bioresource Technol 2004, 93, 9198. [73] Lewandowski, Z., Beyenal, H., Stookey, D. Water Sci Technol 2004, 49, 359-364. [74] Lewandowski, Z., Beyenal, H., Myers, J., Stookey, D. Water Sci Technol 2007, 55, 429-436. [75] Wanner, O., Eberl, H.J., Morgenroth, E., Noguera, D.R., C. Picioreanu, C., Rittmann, B.E, van Loosdrecht, M.C.M. Mathematical Modeling of Biofilms, IWA Task Group on Biofilm Modeling; Publication Date: 01 May 2006. www.iwapublishing.com/ template.cfm?name=isbn1843390876. [76] Caldeira, M., Heald, S.C., Carvalho, M.F., Vasconcelos, I., Bull, A.T., Castro, P.M.L. Appl Мicrobiol Biotechnol 2001, 57, 419-426. [77] De Wildeman, S., Nollet, H., Van Langenhove, H., Verstraete, W. Advan Environ Res 2001, 6, 17-27. [78] Williamson, K., McCarty, P.L. J Water Pollut Control Fed 1976, 48, 9-24. [79] Uemoto, H., Saiki, H. Appl Environ Microbiol 2000, 66, 816–819. [80] Zhang, T.C., Bishop, P.L. Water Sci Technol 1994, 29, 335-344. [81] Lawrence, J.R., Wolfaardt, G.M. Korber, D.R. Appl Environ Microb 1994, 60, 11661173. [82] De Beer, D., van den Heuvel, J.C. Anal Chim Acta 1988, 288, 259-265. [83] De Beer, D., Stoodley, P. Lewandowski, Z. Biotechnol Bioeng 1997, 53, 151-158. [84] Silyn-Roberts, G., Lewis, G. Water Sci Technol 1997, 36, 117-124. [85] Herzberg, M., Dosoretz, C.G., Kuhn, J., Klein, S., Green, M. Water Research 2006, 40, 2704-2712. [86] Kuhn, R.H., Peretti, S.W., Ollis, D.F. Biotechnol Bioeng 1991,38, 340–352. [87] Wanner, O., Gujer, W. Biotechnol Bioeng 1986 28, 314-328. [88] Picioreanu, C., van Loosdrecht, M.C.M, Heijnen, J.J. Biotechnol Bioeng 2001, 72, 205-218. [89] van Loosdrecht, M.C.M., Heijnen, J.J., Eberl, H., Kreft, J., Picioreanu, C. Antonie van Leeuwenhoek 2002, 81, 245-256. [90] Mudliar, S., Banerjee, S., Vaidya, A., Devotta, S. Bioresource Technol 2007 Sep 13. [91] Chen B., Li, Y. Proc. of the 1999 Conference on Hazardous Waste Research, pp.215226; www.p2pays.org/ref/15/14727.pdf. [92] Irvine, R.L., Ketchum, L.H. Jr., Wilderer, P.A., Montemagno, C.D. 1992 United States Patent 5126050. [93] Wilcox, D.P., Chang, E.; Dickson, K.L., Johansson, K.R. Appl Environ Microbiol 1983, 46, 406-416. [94] Grieves, C.G., Stenstrom, M.K., Grutsch, J.F., Walk, J.D. Industrial Wastes, 1977, July/August, 30-35. [95] Hockenbury, M.R., Grady, C P. J Water Pollut Control Feder 1977, 49, 768-777. [96] Hoffman, D.C. Plating, 1973, 60, 157-164.
Immobilized Microbial Cells - Applications and Mass Transfer Phenomena
303
[97] Aizpuru, A., Malhautier, L., Roux, J.C., Fanlo, J.L. Biotechnol Bioeng 2003, 83, 47988. [98] Erhardt, H.M., Rehm, H-J. Appl Microbiol Biotechnol 1989, 30, 312-317. [99] Armenante, P.M., Colela, L.S., Kafkewitz, D., Larkin, M.J. Appl Microbiol Biotechnol 1996, 46, 667-672. [100] Herzberg, M., Dosoretz, C.G., Tarre, S., Green, M. Environ Sci Technol 2003, 37, 4274-4280. [101] Sutherland, J., Adams, C., Kekobad, J. Water Research, 2004, 38, 193-205. [102] Knezovich, J.P., Daniels,J.L., Stenstrom, M.K., Heilmann, H.M. Proc Luxembourg International Symposium on the Rehabilitation of Former Military Sites and Demilitarization of Explosive Ordnance. Nov. 12-16, 1994, pp. 193-202. [103] Lorbeer, H., Starke, S., Gozan, M., Tiehm, A., Werner, P. Water Air Soil Pollut 2002, 2, 183-193. [104] Tiehm, A., Gozan, M., Muller, A., Schell, H., Lorbeer, H., Werner, P. Water Sci Technol 2002, 2, 51-58. [105] Stucki, G., Thuer, M. Appl Microbiol Biotechnol 1994, 42, 167-172. [106] Stucki, G., Thuer, M., Env Sci Technol 1995, 29, 2339-2345. [107] Bouwer E.J., McCarty P.L. Environ Sci Technol. 1982, 16, 836-843 [108] Annadurai, G., Juang, R.-S., Lee, D.-J. J Environ Sci Health Part A 2002, 37, 11331146. [109] Chao, W., Yi, L. Biotechnol Lett 2007, 29, 1353-1356. [110] Caldeira, M., Heald, S.C., Carvalho, M.F., Vasconcelos, I., Bull, A.T., Castro P.M.L. Appl Microbiol Biotechnol 1999; 52, 722-729. [111] Carvalho, M.F., Vasconcelos, I., Bull, A., Castro, P.M.L. Appl Microbiol Biotechnol 2001, 57, 419-426. [112] .Kosseva, M.R., Beschkov, V.N., Peres, C., Agathos, S.N. Bulg Chem Commun 1999; 31: 45-50. [113] Mason, A., Ward, G., Abu-Salah, K., Keren, O., Dosoretz, C.G. Bioprocess Eng 2000, 23, 331-336. [114] Hung,Y.-T., Lo, H.H., Wang, L. K., Taricska, J.H., Li, K.H. In Physicochemical Treatment Processes; Book series “Handbook of Environmental Engineering”; Humana Press ; 2005; Vol. 3, pp. 573-633. [115] Erlanson, B.C., Dvorak, B.I., Speitel, G.E., Lawler, D.F., J Environ Eng 1997, 123, 469-478. [116] Liang, C.H., Chiang, P.-C., Chang, E.-E. Water Research 2007, 41, 3241-3250. [117] Liang, C.H., Chiang, P.-C. J Hazard Mater, 2007, 139, 316-322. [118] Mileva, A., Sapundzhiev, Ts., Beschkov, V. Bioprocess Biosystem Eng 2008, 31, 7585. [119] Kobayashi, F., Nakamura, Y. Biochem Eng J 2004, 21, 93-101. [120] Chen, C.Y., Kao C.M., Chen, S.C. Water Sci Technol 2007, 56, 99-107. [121] Chen, C.Y., Kao, C.M., Chen, S.C. Chemosphere 2008, 1, 133-139. [122] Moser, A. In Biotechnology; Rehm, H-J., Reed, G.; Eds.; Meinheim-Deerfeld Beach, Fa, 1984; vol. 8, pp. 359-363. [123] Moo-Young, M., Kobayashi, T. Canad J Chem Eng 1972 ,50, 162-167.
304
Venko Beschkov
[124] Engasser, J-M., Horvath, C. In Applied Biochemistry and Bioengineering; Wingard, L.B., Katchalski-Katzir, E., Goldstein, L.; Eds.; Academic Press; New York, NY, 1976, vol. 1, pp. 127-220. [125] Stewart, P.S., Robertson, C.R. Appl Env Microbiol 1988, 54, 2464-2471. [126] Dursun, A.Y., Tepe, O. J Hazard Mater 2005,126, 105-111. [127] Dembczynski, R., Jankowski, T., Enzyme Microb Technol 2002, 31, 111-115. [128] Dias, J.C.T., Rezende, R.P., Linardi, V.R. Appl Microbiol Biotechnol 2001, 56, 757761. [129] Harms, H., Zehnder, A.J. Appl Environ Microbiol 1994, 60, 2736-2745. [130] Wijffels, R.H., Englund, G., Hunik, J.H., Leenen, E.J.T.M., Bakketun, A., Guenther, A., Obon de Castro, J.M., Tramper, J. Biotechnol Bioeng 1995 45 1-9. [131] Shreve, G.S., Vogel, T.M. Biotechnol Bioeng 1993, 41, 370-379 [132] Radovich, J.M. Enzyme Microb Technol 1985, 7, 2-10. [133] De Backer, L., Devleminck, S., Willaert, R., Baron, R.G. Biotechnol Bioeng 1992, 322 – 328. [134] Wijffels, R.H., De Gooijer, C.D., Tramper, J. In Immobilised living cell systems: modelling and experimental methods; Willaert, R.G, Baron, G.V., De Backer, L.; Eds.; J. Wiley & Sons, Chichester, 1996; pp. 215-236. [135] Nagashima, M., Azuma, M., Noguchi, S., Inuzuka, K., Samajima, A. Biotechnol Bioeng 1984, 26 992-997. [136] Gosmann, B.; Rehm, H-J. Appl Microbiol Biotechnol 1988, 29, 554-559 [137] Lefebvre, J., Vincent, J.-C. Enzyme Microb Technol 1995, 17, 276-284. [138] Monbouquette, H.G., Sayles, G.D., Ollis, D.F. Biotechnol Bioeng 1990, 35, 609-629. [139] Lefebvre J., Vincent J.-C. Enzyme Microb Technol, 1997, 20, 536-543. [140] Vogelsang, C., Schramm, A., Picioreanu, C., van Loosdrecht,M.C.M., Østgaard, K. Hydrobiologia 2002, 469, 165–178. [141] Schepers, A.W., Thibault, J., Lacroix, C. Enzyme Microb Technol 2006, 38, 324–337. [142] Axelsson, A., Persson, B. Appl Biochem Biotechnol 1988, 18, 231-250. [143] Westrin, B.A., Axelsson A. Biotechnol Bioeng, 1991, 38, 439-446. [144] Yankov, D., Enzyme Microb Technol 2004, 34, 603-610. [145] Killie, K., Hafsjod B., Borgen O., Ratkjie S.K., Hovde, E. AIChE J. 1991, 37, 142-146. [146] Mackie, G.S., Meares, P. Proc Roy Soc London A 1955 232, 498-509. [147] Mavituna, F., Park, J.M. Chem Eng J 1987, 34, Bl - B5. [148] Gutenwik, J. Nilsson, B., Axelsson, A. Biotechnol Bioeng 2002, 79, 664-673. [149] Banerjee, I., Modak, J.M., Bandopadhyay, K., Das, D., Maiti, B.R. J Biotechnol, 2001, 87, 211–223. [150] Wang, H, Seki, M, Furusaki, S. Biotechnol Prog. 1995, 11, 558-564. [151] Monbouquette H.G., Ollis, D.F. Ann. New York Acad Sci 1986, 469, 230–244. [152] Vincent, J.C., Lefebvre, J. J Materials Sci Materials Medicine 1991 2, 234-237. [153] Laca, A., Quiros, C., Garcia, L.A., Diaz, M. Biochem Eng J 1998, 1, 225 – 232. [154] Beschkov, V., Velizarov, S. Bioprocess Eng 2000, 22, 233-241. [155] Cachon, R., Lacroix, C., Divies, D. 1997, 11, 251-256. [156] Obradovic, B., Bugarski, B., Teodosjievic, Z., Nedovic, V., Bugarski, D., VujnjakNovakovic, G. Materials Sci Forum 2005, 494, 531-536.
Immobilized Microbial Cells - Applications and Mass Transfer Phenomena
305
[157] Melick, M.R., Karim, M.N., Linden, J.C., Dale, B.E., Mihaltz, P. Biotechnol Bioeng 1987, 29, 370 – 382. [158] Coronado, C., Botello, J.E., Herrera,F. Biotechnol Prog 2001, 17, 669 -675. [159] Lacroix C., Grattepanche F., Doleyres Y., Bergmaier D. In Application of cell immobilization biotechnology. Focus on biotechnology. [160] Cachon, R., Antérieux, P., Diviès C. J Biotechnol, 1998, 63, 211–218. [161] Krishnan, S., Gowda, M.C., Misra, M.C., Karanth, N.G. Food Biotechnol 2001, 15, 193–202. [162] Doleyres, Y., Fliss, I., Lacroix, C. J Appl Microbiol 2004, 97, 527–539. [163] Shreve, G.S., Vogel,T.M. Biotechnol Bioeng 1993, 41, 370-379. [164] Galazzo, J.L., Bailey, J.E. Biotechnol Bioeng 1990, 36, 417-426. [165] Beschkov, V., Velizarov, S., Georgieva, T. Bulg Chem Commun 1999, 31, 521-535. [166] Petrov, K.K., Yankov, D.S., Beschkov, V.N. World J Microbiol Biotechnol 2006, 22, 337-345. [167] Kosseva, M., Beschkov, V., Pilafova, E. Bulg Chem Commun 1995, 28, 690-702. [168] Burhan, N., Sapundzhiev, Ts., Beschkov, V. Biocheml Eng J 2007, 35, 114-119. [169] Bergmaier, D., Champagne, C.P., Lacroix, C. J Appl Microbiol 2003, 95, 1049-1057.
In: Biocatalysis Research Progress Editors: F. H. Romano, A. Russo
ISBN: 978-1-60456-619-2 © 2008 Nova Science Publishers, Inc.
Chapter XIII
Application of Whole-cell Biocatalysis in Chemoenzymatic, Asymmetric Synthesis of Medically Important Compounds Ewa Żymańczyk-Duda and Paweł Kafarski Departament of Bioorganic Chemistry, Faculty of Chemistry, Wrocław University of Technology, Wrocław, Poland
Abstract Chirality is, in the most cases, the key factor in the safety and efficacy of many drug products [1,2]. Usually only one enantiomer is responsible for the desired activity, whereas its counterpart could be inactive, possess some activity of interest, be an antagonist of the active enantiomer or have a separate activity that could be either desirable or undesirable. Only in a few cases specific compositions of a racemate or an enantiomeric pair demonstrated a synergistic effect. In past decades the pharmacopoeia was dominated by racemates, but since 1980 number of chiral drugs introduced to market have grown markedly. Chiral compounds currently account for at least 50% of sales with the annual sales of single-enantiomer drugs exceeding 150 billions of dollars in 2002 [1,2]. These compounds now represent one-third of all drug sales worldwide. Thus, it is not surprising that chiral intermediates and fine chemicals are in high demand, both from the pharmaceutical and agrochemical industries. During the drug development process, the question invariably arises of which step and which method to choose for introducing chirality. Thus, racemates are usually produced by parallel synthesis for drug candidates - it makes sense to resolve the racemate for the initial animal studies, whereas an optimized, enantioselective synthesis is employed in the course of production. While enantiomerically selective organic synthesis is the traditional approach, using enzymes and enzyme-containing microorganisms to biocatalyze a
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Ewa Żymańczyk-Duda and Paweł Kafarski reaction is becoming increasingly important. However, only 85% of products obtained by biocatalytic processes are enantiomerically pure and in 50% of these processes enantiomerically active substrates are used [3]. This shows that induction of asymmetry is not always the most important feature of industrial biocatalysis. Quite successfully it can be used to carry out conversions that would otherwise require difficult or multiple synthetic chemistry steps. In such cases, biocatalysis can be the preferred route even if chirality is not desired. Moreover, over 25% processes base on kinetic resolution, reaction which affords the desired products with maximal yield of 50% and thus seems to be not interesting from industrial point of view. This is because the second enantiomer is either useful (although in different processes) or is isomerized and processing back as subtrate. The microbial biocatalysts demonstrate a wide variation of activities between genera, within genera and even within species. The range of substrates transformed and the interand intra-species differences in specificity of the individual biocatalysts suggests, that it is possible to provide multiple catalytic agents, especially when the traditional organic process fails or is to expensive to perform. Thus, biocatalysis have become an attractive alternative to conventional catalysts in numerous industrial processes. When compared to chemical catalysis biocatalysts are exquisitely selective and highly precise due to: their substrate selectivity, which allowed distinguishing and acting on the subset of compounds within a larger group of chemically related compounds; their stereoselectivity - the ability to act on a single enantiomer or diastereoisomer selectively and their regioselectivity – ability to recognize one location in a molecule and finally because of their selectivity towards defined functional group in a presence of other equally reactive or more reactive ones. Furthermore, biocatalysts are able to carry out bioconversions under mild conditions – another benefit of using them as industrial catalysts [3-5]. Biocatalysis in drug reseach and production is usually used in multistep processes in concert with more traditional production techniques. Thus, biocatalysts (as both isolated enzyme and whole-cell systems) are increasingly being used to assist in synthetic routes to complex molecules of industrial interest, in so called chemoenzymatic processes [6,7]. Whole-cell biocatalysis is a useful alternative to the use of pure enzymes. Employing whole – cells is a strategy which allowed overcoming many limitation of the enzymatic biotransformations and, what is important, is usually cheaper than using purified enzymes. Additionally, microbial cells used as a catalysts, have their own cofactor regeneration systems, offer wide range of enzymatic activities towards a number of nonphysiological substrates and moreover, usually there are no side reactions except the expected ones [7,8]. Moreover, there is no doubt that, the advantage of microbial biotransformations is the possibility to induce enzymes of defined, desired activity even if they are not constitutively presented inside the microbe cells. This enlarge the offer of possible reactions to be carried out and in fact there exists an enzyme for almost every type of chemical reaction.
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1. Microbial Reductions and Oxidations The usefulness of biotransformations based on the reduction reactions for the introduction of chiral centers into pharmaceutical compounds is a technique, very often used predominantly, but of course the final product is always obtained as an effect of the chemoenzymatic synthesis. Reduction strategies, already applied by the pharmaceutical industry in order to obtain the asymmetric intermediates for further applications, include following reactions: reduction of the carbonyl compounds, reductive aminations, reductions of sulfoxides and alkene double bonds [9]. These reactions are catalyzed by intracellular enzymes aided by the reduced form of certain cofactors. Reduction of the carbonyl compounds is the common, and known for years, way for the introduction the chirality into the prochiral substance. Particular enantiomers of the alcohols serve as a chiral synthons for the next steps of the asymmetric chemical synthesis. There is a huge number of microorganisms which are able to carry out the enantioselective hydrogen transfer from the suitable donor, which obviously in this case, is reducted form of the proper cofactor of the enzyme involved in particular reaction. It is worth to note, that in one living cell there is usually a number of dehydrogenases present and among them there are enzymes of opposite enantioselectivities. It is taken as an advantage, since there is a possibility to control the reaction course by the change of the bioreaction conditions, by the use of suitable biocatalysts or, finally, by modifications of microbial cells. Particular procedures invented for every process often allowed synthesis of pure enantiomers of desired absolute configurations from the same substrate. Enantioselective reduction of carbonyl functionalities has been applied in wide range of chemoenzymatic synthesis and have yielded a number of structurally diverse products. Enzymes that are used in whole-cell bioreductions are often more stable due to the presence of their natural environment inside the cell. Because reductase-catalyzed reactions are dependent on cofactors, one major task in process development is to provide an effective method for regeneration of the consumed cofactors. Many whole-cell biocatalysts offer their internal cofactor regeneration that can be used by adding co-substrates, glucose or, in the case of cyanobacteria, simply light. Chiral β-hydroxy esters are versatile intermediates in the preparation of the series of the various medical agents, including important group of anticholesterol pharmaceuticals. The application of the whole cells of Geotrichum candidum allowed to synthesize methyl (S)-4chloro-3-hydroxybutanoate from the corresponding ketone. This is a key step in the total synthesis of HMG-CoA reductase inhibitor – a cholesterol antagonist affecting the lipid level in blood [10,11] (Scheme 1). The yield and the enantioselectivity of the bioconversion were excellent (up to 98%), what was achieved with the biocatalysts cells cultivated with the supplementation of the glucose or glycerol following the heat treatment of the cells in order to eliminate non-wanted dehydrogenases. The formation of the other HMG-CoA reductase inhibitor, requires the intermediate with two stereogenic centre. This is a diol – (3R, 5S)-dihydroxy-6-(benzyloxy)hexanoate, a product of the Acinetobacter calcoaceticus catalyzed biotransformation of the suitable ketone [12] (Scheme 2). Reaction proceeds through two intermediate monohydroxy compounds, which are reduced upon prolongation of the process. The whole process is easy to perform
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although it requires the addition of glucose and NAD+ as a support for the cofactor regeneration system. A reaction yield of 92% and enantiomeric purity of 98% were achieved when reaction was carried out at 10g/L in a 1L batch.
O Cl
COOCH3
Geotrichum candidum
OH Cl
COOCH3
F HO O OH P
COOCH3
Inhibitor of HMG-CoA reductase Scheme 1.
F OH O
O COOCH3
O
Acinetobacter calcoaceticus
O
O F
N N N N
Inhiibitor of HMG-CoA reductase
OH COOCH3
O
OH OH
+ O
OH O O
O
COOCH3
COOCH3
Scheme 2.
Denopamine – (R)-(-)-α-[3,4-dimethoxyphenylethyl)-amino] methyl-4-hydroxybenzyl alcohol is a β1 adrenoreceptor agonist and coronary vasodilator. It is used in the treatment of congestive heart failure by increasing cardiac pumping function without inducing a significant increase in heart rate. Synthesis of this drug requires the enantioselective
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reduction of the starting ketone and the use of baker’s yeast appeared to be of choice here (Scheme 3) [13]. O OH baker's COOCH3 COOCH3 yeast HO
HO
OH
H N
OCH3
HO
OCH3 denopamine
Scheme 3.
Similarly as denopamine another β-adrenoreceptor agonist, namely selmesterol is also synthesized via innovative, recently developed method, which relays on carbonyl functionality reduction by the cells of Rhodotorula rubra using an anionic surfactant – lauryl sulphate, as a reaction medium [14]. The surfactant may function by establishing micelles that sequester the substrate and microbe [15]. Another microbial stereoselective reduction allowed to obtain the next β-adrenergic receptor antagonists, potentially useful drug against hyperglycemia, intestinal motility disorder or obstructive lung diseases [16]. Initial screening of a number of microorganisms allowed to isolate the strain of the Zygosaccharomyces bailii, which demonstrates the best enantioselectivity against the starting ketone (Scheme 4). The cells of Zygosaccharomysces bailii effectively produced the (R) – chlorohydrin derivative from the corresponding 2chloro-1-[6-(2, 5-dimethyl-pyrrol-1-yl)-pyridin-3-yl]-ethanone simplifying the whole synthesis [16].
N N
N
Zygosaccharomyces bailii
Cl
N
Cl OH
O
N O HOOC
N H
OH
antagonist of adrenergic receptor
Scheme 4.
NH2
Ewa Żymańczyk-Duda and Paweł Kafarski
312 H3CO
O O
COOR
baker's yeast
H3CO
OH O
COOR
OH Scheme 5
NH O
Scheme 5.
Enantioselective reduction was also successfully applied for the production of the synthetic β-lactam antibiotics [17] (Scheme 5). Reaction biocatalyzed by baker’s yeast begins the multistep procedure, which finally resulted in chiral lactam ring, which can serve as a starting substrate for the further chemical modifications. Bioreduction lasting 20h yielded optically pure alcohol of R-configuration. OH
O COOEt
Aurebasidium pullulans
COOEt
pig liver esterase OH
O COOH
Candida
COOH
OH
O
COOCH3
retinoic receptor antagonist Scheme 6.
Application of Whole-cell Biocatalysis in Chemoenzymatic…
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Microbial conversion of the carbonyl functionalities is also a valuable tool for the chemoenzymatic production of chiral retinoids (vitamin A derivatives). The application of the cells of Aureobasidium pullulans resulted in the bioconversion of ethyl 2-oxo-2-(1’,2’,3’,4’tetrahydro-1’,1’,4’,4’-tetramethyl-6naphthalenyl)acetate into corresponding alcohol of Rconfiguration, which is then converted to retinoid receptor γ-specific agonist [18], which is medically used as dermatological and anticancer drug. This reaction was previously catalyzed by the other microbes, giving satisfactory enantiomeric purities but the chemical yields of the process were far too low. The application of A. pullulans allowed to reach 98% yield of the process (Scheme 6). Hydrolysis of the obtained ester with pig liver esterase afforded the corresponding acid with good yield. The same acid could be obtained by direct reduction of 2-oxo-2-(1’,2’,3’,4’-tetrahydro-1’,1’,4’,4’-tetramethyl-6naphthalenyl)acetic acid with Candida utilis or Candida maltosa (Scheme 6). Application of baker’s yeast bioreduction also allowed to obtain (R)-fenfluramine, a drug that was part of the Fen-Phen anti-obesity medication. Fenfluramine causes the release of serotonin by disrupting vesicular storage of the neurotransmitter, and reversing serotonin transporter function. The end result is a feeling of fullness and loss of appetite. However, the drug was withdrawn from the U.S. market in 1997 after reports of heart valve disease and pulmonary hypertension, including a condition known as cardiac fibrosis. When being marketed the key step of its synthesis relayed on biocatalytic reduction of corresponding ketone (Scheme 7) [19]. The other example of the stereoselective reduction catalyzed also by baker’s yeast cells is production of (1R)-3-chloro-1-phenylpropan-1-ol, a crucial step in the Prozac (fluoxetine hydrochloride) synthesis (Scheme 8). Prozac is an effective antidepressant and is active against a wide range of symptoms such as anxiety, alcoholism, chronic pain and eating disorders [20,21]. Similarly as all the examples of baker’s yeast reductions this process illustrates the simplicity and the effectiveness of this procedure, which resulted in enantiomerically pure product with excellent chemical yield. The alternative procedure for production of Prozac and related drugs, such as atomoxetine and nisoxetine is based on the reduction of the prochiral 3-oxo-3phenylpropanenitrile. In this case baker’s yeast cells are uneffective because alkylation predominates over biocatalytic reduction yielding undesired side-products. baker's yeast
F3C
F3C OH
O
F3C HN fenfluramine Scheme 7.
Ewa Żymańczyk-Duda and Paweł Kafarski
314 O
Cl
OH
baker's yeast
Cl
CF3 O N H
Prozac
Scheme 8.
O CN
recombinant Escherichia coli
OH CN
CF3
H3CO O
O
O N H
atomoxetine
N H
N H nisoxetine
Prozac
Scheme 9.
The application of the recombinant Escherichia coli, in which yeast reductase was overexpressed allowed to overcome the problem of low yield of reduction. Similarly, the same reductase was overexpressed in eukaryotic microbe - Curvularia lunata [22]. Although being significantly more expensive, than the simple use of cheap baker’s yeast, the use of recombinant microorganisms is more universal because it is leading to the series of structurally related chiral drugs when starting from common chiral substrate (Scheme 9). Chlorhydrine derivatives with two stereogenic centers of (1S,2S) or (1S,2R) configurations are essential for the synthesis of two HIV protease inhibitors, namely popular aza-peptidomimetic Atazanavir and clinically tested BMS-186318. (1S,2S)-[3-Chloro-2hydroxy-1-(phenylmethyl)propyl]carbamic acid, which is precursor of Atazanavir [23,24] is produced by Rhodococcus erythropolis enantioselective reduction of the corresponding 2-oxo compound, whereas the isomer of (1S,2S) configuration being a chiral building block for BMS-186318 formation is obtained as a result of the bioconversion of the same substrate catalyzed by Streptomyces nosdus [25,26]. The single stage process resulted in pure asymmetric products, which formed with 80% yield (Scheme 10).
Application of Whole-cell Biocatalysis in Chemoenzymatic…
315
O O
N H
Cl O
Streptomyces nosdus
Rhodococcus erythropolis
O O
O N H
Cl
O
OH
N H
Cl OH
O N
O N
O
O O O
O N H
OH
N H
OH
BMS 186318
N H
O
H3CO
OH
O N
N H
H N
COOCH3
Atazanavir
Scheme 10.
Diltiazem hydrochloride is a benzothiazepinone calcium channel blocker, which is used as an antianginal and antiperhensive agent. The vital step of its production is a bioreduction catalyzed either by baker’s yeast or Nocardia salmonicolor [27]. This multistep chemoenzymatic synthesis is a typical example of the role of biocatalysis in drug production (Scheme 11). In same cases specific reductases have to be used in order to achieve the goal. A good example is the use plant troponine reductase I from Datura stramonium to produce squalene synthase inhibitor used for the treatment of arteriosclerosis or bronchodilators. This compounds as well as its analogues is produced from one crucial chiral intermediate – (R)-3quinuclindol, which is obtained from the corresponding starting ketone [28] (Scheme 12). Since the use of plant enzyme did not appear give satisfactory results the reaction is catalyzed by the recombinant Escherichia coli. Quite unusual is enantioselective microbial reduction of hydroperoxides. When using topsoil isolate of Bacillus subtilis various hydroperoxides were reduced yielding structurally variable (R)-hydroperoxides (see representative example in Scheme 13) with moderate to good enantioselectivities. In contrast to the bacteria, the fungus Aspergillus niger displayed the reverse sense of enantioselectivity [29].
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316
OCH3
S
OCH3
biocatalyst
S
O N H
OH N H
O
O
OCH3
S O N
Scheme 11
N
O O Dilitazem
Scheme 11.
Whole-cells reduction of sulfoxides was also employed for the resolution of the racemic sulfoxides [30,31]. Thus, the strains of Escherichia coli or Proteus vulgaris cultivated anaerobically are able to reduce racemic omeprazole. Final, enentiomeric product is a proton pump inhibitor used in the treatment of dyspepsia, peptic ulcer disease, and gastroesophageal reflux disease (Scheme 14). Reductive amination is a chemical reaction which involves the conversion of a carbonyl group to an amine. In the case of this reaction microorganisms are preferentially used because isolated enzymes are too expensive. A good example is synthesis of (S)-2-amino-5-(1,3dioxolan-2-yl)-pentanoic acid (lysine ethylene acetal), a key intermediate in the synthesis of Omapatrilat, a vasopeptidase inhibitor and thus potent antihypertensive drug. This particular example is very interesting because it illustrates the cooperation of two microorganisms Escherichia coli and Candida boidinii, for the reductive ketone amination [32,33] (Scheme 15). It is worth to note that in this case one from the microbes serves as a cofactor regenerating system (recombinant Escherichia coli cells) whereas the second one acts as a real biocatalyst. Similar results were achieved by the use of recombinant Escherichia coli or Pichia pastoris, in which phenylalanine dehydrogenase from Trigonopsis intermedius was overexpressed [34].
Application of Whole-cell Biocatalysis in Chemoenzymatic… H O
H
O
OH
N
N
317
O
N
HO
S
inhibitor of squalene synthase
Scheme 12.
OOH
Bacillus subtilis or horsedish cells
Aspergillus niger
OOH
OH
OOH
+
OH
+
Scheme 13.
H N
OCH3
O S
N
H3CO
N Proteus vulgaris
H N S H3CO
+
N N (+)-omeprazole
Scheme 14.
H N
OCH3
O
OCH3 S
H3CO
N N
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NH2
Candida boidinii
O
COOH O
O
Escherichia coli
COOH O
S H O HS
N N H
O
COOH
Omapatrilat Scheme 15.
O O
O
baker's yeast O
(3R)-cryptoxanthin Scheme 16.
There are a few literature reports about the application of the stereoselective double bond saturations using whole-cells biocatalysts in a pharmaceutical industry. However, there is a group of compounds – retinoids, which are formed via this particular microbial reduction. These chemicals are also prepared in a different although reductive manner – by carbonyl functionalities biotransformation. Here, baker’s yeast cells are employed for the reduction of the starting oxoisophorone to (6R)-2,2,6-trimethylcyclohexane-1,4-dione, which is then converted into (3R)-cryptoxanthin – vitamin A precursor [35] (Scheme 16). Microbial oxidations are less commonly applied than microbial reductions. This approach, similarly as in the case of the reduction, employs intracellular enzymes belonging to oxydo-reductases. The oxidative attempt allowed implementing chirality via the oxidation leading to the asymmetric epoxides or by the introducing one or two oxygen atoms into the starting substrates by hydroxylations or diooxygenations. Microorganisms screened for the particular oxidative procedure, are mostly active only towards structurally similar substrates.
Application of Whole-cell Biocatalysis in Chemoenzymatic…
319
OH
O
O
O N R
Sphingomonas
NH N R
O
N
O
O
O HO
O carbapenem
Scheme 17.
OH
OH OH
Rhodococcus
Rhodococcus
N
OH
N
OH
H N
N
Scheme 18
N H
O
OH
O
Crixivan
Scheme 18.
Microbial hydroxylations are predominantly used for the regioselective introduction of hydroxy group into the wide range of substrates. However, there are also some examples of the chiral centre creation via microbial mono- or diooxygenations. Hydroxylation of Nprotected 2-pyrrolidinones by resting cells of various Sphingomonas allowed to obtain both enantiomers of 4-hydroxy-2-pyrrolidinone (Scheme 17). The presence of glucose, which contributed to cofactor regeneration, was mandatory. The (S) isomer is a key intermediate for the synthesis of the carbapenem antibiotic CS-834 [36] and the nootropic drug – (S)– oxiracetam [37] whereas the (R)-isomer is useful as a substrate for the production of antidepressant agent – (R)–rolipram [38], anticonvulsant (R)-γ-amino-β-hydroxybutyric acid and the antihyperlipoproteinemic L-carnitine [39,40]. The formation of the intermediate in the synthesis of potent HIV protease inhibitor Crixivan was achieved by using various isolates of Rhodococcus (Scheme 18). Depending on isolate biocatalytic transformation of indene leads either to cis-(1S,2R)-indandiol or to trans(1R,2R)-indandiol. The structure of the product resulted from the predominant activity of either monooxygenases or dioxygenases. Thus, the isolates were able to implement one as two hydroxyl group into the starting compound. The yield of the reaction was limited by the
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320
indene toxicity, which has to be carefully monitored during the process [41] in order to allow maintenance of viability of the cells. Tiamulin is diterpene antibacterial agent with a pleuromutulin (isolated from the basidiomycetes strain – Pleurotus) chemical structure. It is used in veterinary medicine for treatment of prophylaxis of dysentery, pneumonia and mycoplasmal infections in pigs and poultry. Its oxidation catalyzed using Cuninghamella echinualta or Streptomyces griseus [42], most likely by cytochrome P450, yielded promising antiinfective agent (Scheme 19), thus showing that the detoxification of pleuromutulin did not result in its deactivation. Epoxidations are also valuable tool for the synthesis of chiral compounds. This strategy was successfully applied for the synthesis of the key intermediate in the β-blockers production by enenatioselective reaction of prochiral aryl allyl ethers. Psudomonas oleovorans was found to be microorganism of choice and (S)-aryl glycidyl ethers were obtained (Scheme 20). They are precursors of (S)-metoprolol and (S)-atenolol – effective agents against angina pectoris, cardiac arrhythmias and migraine [43,44]. Stereoselective epoxidation was also applied as a key step in the synthesis of the potassium channel openers, which play a major role in neuronal excitability. The Mortierella ramanniana cells were applied as a biocatalyst for oxidation of the 6-cyano-2,2-dimethyl-2H1-benzopyran, which resulted in chiral epoxide. This epoxide was subsequently converted by means of few chemical reactions into the new flavonic channel opener [45] (Scheme 21). OH
OH Cuninghamella ehinulata
HO
OH
O
OH
O O
O
pleuromutulin Scheme 19. Pseudomonas oleovorans
R
R
O
O
O
H2N O
O OH atenolol
Scheme 20.
O
N H
O OH metopropol
N H
Application of Whole-cell Biocatalysis in Chemoenzymatic… Mortierella ramanniana
NC
321
O
NC O
O
N
O OH
NC O
Scheme 21 Scheme 21.
Quite unusual is oxidation of (exo,exo)-7-oxabicyclo[2.2.1] heptane-2,3-dimethanol accompanied with formation of corresponding (S)-lactol and (S)-lactone (Scheme 22). This is the crucial step in the total synthesis of the thromboxane A2 antagonist, used to treat vascular disorders. The reaction is effectively catalyzed by whole cells either Nocardia globurula or Rhodococcus sp. [46].
O
OH OH
OH
Nocardia globurula
O
O
O
+
O
O
COOH O O HN O
N H thromboxane A2 antagonist
Scheme 22.
2. Hydrolytic Organisms Hydrolases are the most commonly used in biocatalytic processes and constitute over 75% of commercially available enzymes with lipases being the most important owing to their application in a wide variety of processes. They are efficient catalysts, which enable a
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322
number of esterifications, transesterifications and hydrolysis reactions with high chemo-, regio- and enantioselectivity, accepting a broad range of synthetic substrates with vast industrial importance. Majority of lipases are secreted extracellulary and are readily separable from microbial growth media. Moreover, they belong to serine hydrolases and do not require any cofactor for activity [47-49]. Thus, they are easily available and relatively cheap enzymes, which causes that the use of lipolytic microorganisms as biocatalysts is strongly limited. The only exception is Yarrowia lypolitica, yeast routinely isolated from various food media, which exhibits enormous hydrolytic potency (Scheme 23) [50]. Lipase B from Candida antarctica is the most robust enzyme in organic synthesis. It has a broad range of applications including polymerizations, resolutions of alcohols and amines, modifications of sugars and sugar-related compounds, desymmetrization of complex drug intermediates and ring opening of β −lactams [51]. Although C. antarctica was isolated from the bottom of Antarctic lake Vanda and thus is a member of extremophile family, lipases isolated from that organism are exceptionally thermostable. For industrial purposes these lipases are obtained by using recombinant Aspergillus oryzae, which is far easy to cultivate. There are also rare examples of the use of recombinant cells producing Candida lipases in organic synthesis [52,53]. Chiral α-hydroxy ketones are important synthons for the asymmetric synthesis of natural products. The effective method of their preparation from simple ketones relays on chemoenzymatic reaction, in which chirality is introduced by microbial hydrolysis of αacetoxyketones (Scheme 24). In the case of the most effective strains of Rhizopus oryzae desired products were obtained in 48% yield with very high enantiomeric excess [54]. The impure second enentiomer was isomerized by the action of strong base and recycled into the process. (R)-1-(3-Chlorophenyl)-2-hydroxypropan-1-one, obtained in this manner is a starting material for production of Bupropion marketed for the treatment of depression and approved as an aid to smoking cessation, whereas (R)-1-(2,4-difluorophenyl)-2-hydroxypropan-1-one is a substrate for production of two antifungal compounds being under clinical studies.
OAc
O Yarrowia lipolytica
O R O
AcO
OAc
n Scheme 23.
OAc
Application of Whole-cell Biocatalysis in Chemoenzymatic… O
O
cyclohexane reflux
O
Rhizopus oryzae
Mn(OAc)3 X
323
OAc
X
OH X DBU/hexane-THF
O
O
Cl
Cl OH
HN Bupropion
N
N
N
N
N O
HO
N HO
and
S F
F
OH
F
F
SO2CH3
F
N
F
Scheme 24
CN
Scheme 24.
4-(R)-Hydroxycyclopent-2-en-1-(S)-acetate is an important intermediate in the synthesis of cyclopentenoid natural products such as prostaglandins, prostacyclins and tromboxanes, and some drugs against AIDS. This has resulted in various approaches for its preparation. Promising and simple seems to be the use of Trichosporon beigelii as a catalyst for the enantiomeric hydrolysis of readily available meso-cyclopent-2-en-1,4-diacetate which affords the desired enantiomer of 4-hydroxycyclopent-2-en-1-(S)-acetate (Scheme 25). In this case enhancement of the stereoselectivity of the reaction was reached by the application of ethanol as a co-solvent [55]. O
O Scheme 25.
O
Trichosporon beigelii
O
phospate buffer ethanol
O
O O
OH
OR
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324
1. Prins reaction
COOCH3
H3COOC
+
2. Jones oxidation 3. Esterification
O
O Bacillus coagulans COOCH3
HOOC
HO COOH
+
O HO
OH
O
D-cloprostenol
O
Cl Scheme 26.
O
O O
Bacillus coagulans
O
O HO
O
O
+
O
O
O
Scheme 27.
Another procedure for production of prostaglandin derivatives considers the use of 2oxotricyclo[2.2.1.0]heptan-(R)-7-carboxylic acid, which is synthesized in its racemic form from readily available norbornadiene. Resolution of this acid was achieved by precipitation with chiral amines or by enenatioselective hydrolysis of its methyl ester with commercial lipases, however, both methods are not satisfactory. Application of a selected strain of Bacillus coagulans resulted in desired enatiomer with 50% yield and very high enantioselectivity (Scheme 26). (R)-Acid was then used as a substrate for the preparation of D-cloprostenol (a powerful luteolytic agent approved for use in beef and cattle) by standard procedure [56]. B. coagulans NCIMB 9365 possesses two cell-associated competing esterases: carboxylesterase 1, which is enantioselective towards (R,S)-1,2-O-isopropylidene glycerol benzoate and thermostable at 650C, and carboxylesterase 2, which is not-enantioselective with a maximum activity at 35 0C. Whole cells were, therefore, modified by heat treatment in order to knock-out carboxylesterase 2 selectively and then were applied to resolve (R,S)-1,2O-isopropylidene glycerol benzoate (Scheme 27) [57]. This is a classical example of manipulation of whole-cell specificity. Levofloxacin, a member of fluoroquinolone antibiotics, is used to treat pneumonia, chronic bronchitis and sinus, and urinary tract, kidney, and skin infections. It is synthesized via key intermediate, namely (S)-7,8-difluoro-2,3-dihydro-3-methyl-4H-1,4-benzoxazine (Scheme 28). Chemoenzymatic process of its preparation in optically pure form was achieved by application of certain strain of Bacillus sp. [58], elected by screening over 500 various
Application of Whole-cell Biocatalysis in Chemoenzymatic…
325
microorganisms. Hydrolysis is extremely effective since both enantiomers (amine and unhydrolyzed amide were obtained in 48% yields with 98% ee). Large scale (20 L) synthesis of enantiomerically pure chiral arylalkylamines, being standard pharmaceutical building blocks was elaborated in laboratories of Novartis using whole cell biotransformations with microorganisms containing novel enantioselective amidohydrolases [59]. Although the chosen microorganisms produce stable and highly selective aminohydrolases the use of whole-cell system appeared to be of choice. Thus, the use of (S)-selective Rhodococcus equi and Rhodococcus globerulus and (R)-selective Arthrobacter aurescens afforded desired amines (see representative example in Scheme 29) with excellent enantioselectivity and of high yield (45% after distillation of amine). It is worth to mention that all the three organisms also exhibit promising nitrile hydrolase and epoxide hydrolase activities [60, 61]. F
O
F
N
F
F Bacillus sp.
O
+ F
NH
F
O N O
O
O F
COOH
N N
N O
Scheme 28.
O HN
O Rhodococcus equi
HN
NH2
+ or R. globerulus
O
O HN
Scheme 29.
Arthobacter aurescens
HN
NH2
+
Ewa Żymańczyk-Duda and Paweł Kafarski
326 O
H
Pseudomonas solonacearum
N
HOOC
O
NH2
H N
+
O N HO
N
N
N
H NH2
carbovir Scheme 30.
(±)-2-Azabicyclo[2.2.1]hept-5-en-3-one is a versatile intermediate in the synthesis of carbocyclic nucleotides. The use of Pseudomonas solonacearum enabled to obtain both optical forms of the lactam in very high optical purity (over 98% ee) in rapid, facile and large-scale biotransformation process (Scheme 30). The use of P. solonacearum is superior over the use of purified enzymes such as β–lactamase or savinase [62]. Fusarium oxysporum produces specific lactone hydrolyzing enzyme (lactonase), which is industrially used by Daiichi Fine Chemicals for stereospecific hydrolysis of D-pentalactone [63], an intermediate in the synthesis of folic acid (vitamin B9) via panthotenic acid (Scheme 31). The process is performed with mycelia entrapped in calcium alginate in the presence of calcium chloride. With concentration of the substrate as high as 300 g/l the yield of the reaction is over 40% and the product is obtained with enantiomeric excess higher than 90%. It is worth to note that mycelium retains 70% of the activity after 180 batch reactions. The estimated half-life of the lactonase activity of the immobilized mycelia is 6000 h, which is 35 times higher than that of the free mycelia. 3000 tons of D-pentalactone is produced annually using this process.
OH
OH
OH
COOH O
O
O
OH
O
O HO OH Scheme 31.
N H
COOH panthotenic acid
Application of Whole-cell Biocatalysis in Chemoenzymatic…
OSO3H
Sulfolobus acidocalvarius
OSO3H
327
OH
+
pH 2-3 Scheme 32.
Sulfatases catalyze hydrolytic cleavage of sulfate ester bond. The steric course of this reaction might be controlled by the choice of the appropriate subtype of sulfatase enzyme. Sulfatases offer the possibility of transformation of a racemic secondary alcohol into a single stereoisomeric product in 100% theoretical yield through a kinetic resolution. This can be accomplished because biocatalytic process undergoes with retention of configuration (representative example is shown in Scheme 32). Thus, aerobically grown Sulfolobus acidocalvarius catalyzes hydrolysis of secondary alcohol sulfates in high yield with excellent enantioselectivity [64]. Nitrilases are a group of enzymes that have good potential in the chemical process industry for conversion of nitriles to a wide range of useful products and intermediates. The enzymes are mostly bacterial and there are two main groups: nitrilases and hydratases with the latter being more readily available than the former. Traditional chemical processes for the hydrolysis of nitriles are expensive requiring high temperature, also inefficient as many byproducts are produced, which means that the isolation of the commercial product is more difficult and there is a disposal problem for the unwanted by-products. On the other hand nitriles are readily available by simple organic synthesis. Thus, the use of nitrilases is a method of choice. Unfortunately some nitrilases loss their activity upon exposure to oxygen, most likely because oxygen reacts with their active-site cysteine [65]. Therefore, the use of microbial whole-cells is advantageus. The extensive screening of different degraders has led to the selection of the most efficient organisms, mostly various Rhodococcus strains, capable efficiently biotransform a wide range of nitriles using either a two-step pathway (nitrile hydratase and amidase) or a one-step pathway (nitrilase) and finally yielding the corresponding carboxylic acids and ammonia (Scheme 33) [66].
R CONH2
nitrile hydrolase
H2O
H2O
amidase
R C N nitrilase
H2O NH3 Scheme 33.
NH3
R COOH
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328
Rhodococcus sp.
or
+
CN
CONH2
COOH
CONH2
S HOOC NH2
HN
H
O
O
O
cilastatin
anthoplanone
Rhodococcus sp. R
CN
racemic cis-
phosphate buffer/hexane (1:1) R = Ph
R
+
+ R
COOH
CONH2
32%
COOH R
R
49%
CN
19%
R = CH3
CN
racemic transOCH3
S N curacin A
NH2
tranylcypromine
Scheme 34
Scheme 34.
Thus, Rhodococcus sp. AJ270 containing both sets of enzymes was successfully applied for successful resolution of variously substituted cyclopropanenitriles (Scheme 34), a versatile building blocks for such important drugs as cilastatin (protects antibiotic imiphenem from degradation by renal dehydropeptidases), curacin (a potent cancer cell toxin obtained from strains of the tropical marine cyanobacterium Lyngbya majuscule), anthoplalone (sesquiterpenic anticancer agent produced by Anthopleura pacifica) or quite simple structurally tranylcypromine (strong antidepressant) [67,68]. Molecular recognition of an enzyme toward a substrate is a crucial step in enantiospecific or highly enantioselective biocatalytic transformations. When a non-natural substrate is used, a general practice in synthetic biotransformations, low enantioselectivity, and slow reaction is frequently encountered because of nonspecific interaction between the enzyme and the
Application of Whole-cell Biocatalysis in Chemoenzymatic…
329
substrate. To circumvent these problems, structural modifications of enzymes such as sitedirected mutagenesis and directed evolution have enjoyed the success in upgrading the performance of enzymes. As an alternative to biotechnological approaches, substrate engineering has been shown to be useful in improving the efficiency and selectivity of biocatalysis. Substrate engineering, in contrast to protein engineering, modifies the structure of small organic molecular substrates to best fit into the active site of the enzyme. These approaches are therefore relatively simple, easy to handle, and cost-effective. For example, catalyzed by the Rhodococcus erythropolis whole cell catalyst, the O-benzylated β-hydroxy alkanenitriles underwent remarkably high enantioselective biotransformations, whereas the biotransformations of free β-hydroxy alkanenitriles gave very low enantioselectivity (Scheme 35). Thus, the easy manipulations of O-protection and O-deprotection, excellent chemical and enantiomeric yields of biotransformations, along with the scalability render this enzymatic transformation attractive and practical for the synthesis of highly enantiopure βhydroxy alkanoic acids and their amide derivatives [68]. Enantiomerically pure epoxides and chiral diols are important building blocks in organic synthesis and can be used as key intermediates in the synthesis of more complex enentiopure compounds. Thus, several production methods have been elaborated with the use of epoxide hydrolases being particularly attractive. Aryl epoxides are potentially useful substrates for the synthesis of β-blockers and anti-obesity compounds. Such a procedure is represented by hydrolysis of phenylglycidyl ether with lyophilized whole cells of Bacillus megaterium, which resulted in hydrolysis of R-isomer yielding S-epoxide with good yield and enantioselectivity (Scheme 36) [69]. Styrene oxide was hydrolyzed by Aspergillus niger LCP 521 with an opposite enantiopreference ((R)-selective) and regioselectivity compared to conversion catalysed by Beauveria sulfurescens ATCC 7159 ((S)-selective). Thus, the combined use of both organisms yielded (R)-phenylethanediol in 92% yield and 83% e.e. [70]. Racemic hydantoins are readily available by simple chemical process. The biocatalytic conversion of hydantoins to amino acids has been recognized in recent decades for potential application in the industrial production of amino acids, both natural and non-natural. The utility of non-natural amino acids is well represented by application of L-tert-leucine as a building block for various drugs (Scheme 37). Rhodococcus erythropolis
O
+
O
O COOH
CONH2
CN
Scheme 35.
O
Scheme 36.
O
Bacillus megaterium
O
OH
O
+
O
OH
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330
COOH NH2
OH R
H N
O
H N
N H HO
N H
O
R HO
H N
O
H N
N H
O
CH3
O
antiinflammatory, Hoffmann La Roche antiviraly, Sandoz F O H
O
COOH N H
N CN
N
O O
H N
N H
anticancer, Zeneca O
H N O
O N H
COOH
antivitral, Biomega
Scheme 37.
The biocatalysis involves two consecutive hydrolysis steps, catalysed by hydantoinase and N-carbamoylamino acid amidohydrolase. Hydrolysis is enantioselective and results in desired enantiomer of amino acid and unreacted enantiomer of hydantoin. The latter one is easily racemized by hydantoin racemase. Interesting version of this process is the use of recombinant Eschericha coli obtained by cloning D-hydantoinase and D-carbamoylase from Agrobacterium radiobacter and hydantoon racemase from Agrobacterium tumefaciens [71,72]. After optimizing reaction conditions nearly 100% conversion of D,L-hydantoins into D-amino acids was achieved (Scheme 38). H O
R
O
HN
NH
HCN (NH4)2CO3
racemase
O
D-hydantoinase
O
R D-carabamoylase H2N Scheme 38.
COOH
H2N
R
O
HN
NH
R N H
COOH
+
O
Application of Whole-cell Biocatalysis in Chemoenzymatic…
331
OH HO HO NH2
HO
OH
valienamine
HO HO HO
H3C O N H HO OH O HO acarbose
OH
OH OH
O OH O HO
HO HO
O OH
OH N
HO
OH
OH voglibose OH HO
Scheme 39
HO HO
H3C O N H HO OH OCH3 acarviosin
Scheme 39.
Acarbose (Scheme 39), a strong inhibitor of glucosidase, is an anti-diabetic drug used to treat type 2 diabetes mellitus and, in some countries, prediabetes. It is obtained by fermentation, from secondary metabolic products of soil microorganism – Actinoplanes sp.. However, in broth six other oligosaccharides of related structure are also present. At present they are isolated and discarded in factories. Therefore the way of their utilization is a significant problem. All of these side products contain valienamine as an terminal sugar. This non-typical sugar is, in turn an important substrate in synthesis of other valuable anti-diabetic drugs as acarviosin, valienamine and valiolamine (voglibose) (Scheme 39). Stenotrophomonas maltrophilia, isolated from soils near Hangzhou (Zhejiang, China) appeared to be an organism of choice for hydrolytic degradation of the mentioned sideproducts and the production of valienamine from that waste products [73].
3. Microbial Carbon-carbon Formation The stereoselective carbon-carbon formation is a key topic in today’s synthetic organic chemistry. Enzymes applied in biocatalytic C-C bond forming reactions are aldolases, transaldolases, transketolases, cyclases and hydroxynitrile lyases (oxynitrilases) [74]. Oxynitrilases catalyze the formation and cleavage of cyanohydrins. Cyanohyrins, which are extremely useful blocks in synthesis of many compounds of pharmaceutical interest, are obtained stereospecifically by addition of hydrogen cyanide to aldehydes or ketones. Reactions catalysed by oxynitrilases offer economical access to enantiomerically pure
Ewa Żymańczyk-Duda and Paweł Kafarski
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cyanohydrins, -hydroxy acids, aminoalcohols and other downstream products. The biocatalitic application of almond meal was described for the first time in 1913 [75]. Since cyanohydrin fission plays an important role in nature and is involved in plant defence where hydrogen cyanide is liberated upon plant damage, the plant cells are most often used as bioctalysts, with almond meal being the most popular. Pure enzymes are used rarely since only those from almond (Prunus amygdalis), Hevea brasiliensis and Manihot esculenmta are available in sufficient quantities [76]. The synthesis of (R)-pantolactone was achieved in three-step process consisting of: aldol reaction of 2-methylpropanal with formaldehyde followed by biocatalytic hydrocyanation of the obtained aldehyde to afford (R)-2,4-dihydroxy-3,3-dimethylbutylnitrile, which was subsequently acid hydrolysed to afford (R)-pantolactone (Scheme 1). The vital step of the synthesis is enantioselective asymmetric hydrocyanation catalyzed by almond meal, or either apple or plum kernels. Hydrogen cyanide or acetone cyanohyrdrin were used as second substrate [77]. Enantiomeric aromatic cyanhydrines are important intermediates of pharmaceutical interest. It is well documented in the case of production of (R)-salbutamol and (R)tetrabutaline, potent and short-acting β2-adrenergic receptor agonists used for the relief of bronchospasm in conditions such as asthma and chronic obstructive pulmonary disease. Preparation of enantiomerically pure cyanohydrine is the crucial step of their synthesis [78]. Synthesis of aromatic cyanhydrines might be achieved by application of various crude sources of oxynitrilese such as almond meal [79] and meals obtained from seeds of guanabana (Annona muricata) [80], common vetch (Vicia sativa) [81] (for representative example see Scheme 41).
HCHO
CHO
HO
CHO
almond meal HCN
OH CN
HO
OH O O
Scheme 40 Scheme 40.
OH
O
CN Scheme 41.
Vicia sativa
Annona muricata O
CHO
O
OH CN
Application of Whole-cell Biocatalysis in Chemoenzymatic… O O
O Bacillus cereus
H
+
333
HO
HO O
HO frambinone Scheme 42.
The aldol condensation has been an effective method in organic synthesis for the formation of carbon-carbon bonds and much effort has been directed toward making these reactions proceed in a stereoselective way. Complementary to the chemical methods is the use of aldolases, which in general are known to induce high stereoselectivity [82]. Aldolases are commonly used as isolated enzymes and the reports on the use of whole-cell biocatalysts are very scarce. An exemption is the use of several bacteria possessing 2-deoxyribose-5phosphate aldolase to produce an important component of flavouring and perfume raspberry ketone (frambinone) in the reaction between acetone and 4-hydroxybenzaldehyde (Scheme 42) [83]. Raspberry ketone represents a total market value of between €6 million and €10 million. The possibility of producing its direct precursor through a simple process using bacteria is of considerable interest to the flavour market and the food industry as a whole. Condensation of two readily available compounds - acrolein and pyruvate, catalysed by baker’s yeast afforded valuable (R) -(−)-3-hydroxy-1-penten-4-one with good yield and of high enantiomeric purity [84]. Although the major product is accompanied by small quantities of its tautomer the procedure is economic, simple and very efficient (Scheme 43). The pharmaceutical industry has long been indebted to fermenting yeast for asymmetric carbon-carbon bond synthesis. Using pyruvic acid and benzaldehyde as substrates (R)phenylacetylcarbinol was obtained in a stereoselective manner (Scheme 44). This product treated with methylamine over platinum catalyst under athmosphere of hydrogen afforded Lephedrine [85].
H O
Scheme 43.
baker's yeast
O +
COOH
-CO2
OH
O + OH major
O minor
Ewa Żymańczyk-Duda and Paweł Kafarski
334 O
baker's yeast
O H
+
COOH
O
-CO2 OH CH3NH2 H2/Pt
NHCH3 Scheme 44 OH L-ephedrine Scheme 44.
It is well established that high pressure carbon dioxide has an sterilizing effect upon bacteria. However, there are bacteria able to cope with this gaseous substance and use it as a source of carbon for growth. Thus Bacillus megaterium is able to catalyse the reversible carboxylation of pyrrole to pyrrole-2-carboxylate (Scheme 45). When culturing bacteria under elevated pressure of carbon dioxide (100 barr, 400C) in phosphate buffer (pH 5.5) containing ammonium acetate (nitrogen source for bacterial growth) the yield of pyrrole carboxylate was found to be twelve-fold higher that in atmospheric pressure (1 barr) [86]. Pyrrole-2carboxylic acid is a valuable synthon for preparation of many pharmaceutical products (for representative examples see Scheme 45) NH2 O CO2
N N H
N H
HO
COOH Cl
O
O
antiviral agent Cl
N
O
COOH
O S
N N
COOEt
COOH
N
N NH2 inhibitor of interleukin I
Scheme 45.
antiviral agent
blocker of angiotensin II
O
Application of Whole-cell Biocatalysis in Chemoenzymatic…
Thauera aromatica
OH
CO2
335
OH HOOC
Scheme 46.
Second example of this unusual reaction is carboxylation of phenol by whole cells of Thauera aromatica containing phenylphosphate carboxylase (Scheme 46). Reaction is highly regioselective and carboxylation undergoes selectively in para position. Bacteria do not require carbonate to catalyze carboxylation but is able to use gaseous carbon dioxide directly [87].
4. Miscelanous Processes Resveratrol is a constituent of several food products and medicinal plants. It is a potent phytoalexin, which displays an array of pharmaceutical and antimicrobial activities, however, its in vivo pole is questionable. Piceid, a glycosylated form of resveratrol, acts as prodrug and enable to avoid drawbacks resulted from application of underivatized resveratol. Preparative scale biotransformation of resveratol with whole-cell suspensions of Bacillus cereus resulted in the production of piceid in good yield (Scheme 47) [88]. Ferrulic acid is an abundant and renewable source of aromatic compounds from agricultural byproducts. Among these vinylguaiacol is the most commonly occurring ferrulic acid coversion product. Whole-cells of Bacillus pumilus were used to decarboxylate ferrulic acid (Scheme 48) with good yield when organic solvents such as n-hexane, n –heptane or noctane were used in biphasic system [89].
OH HO HO OH
OH O
OH HO
Bacillus cereus
OH HO
resveratrol Scheme 47.
O
piceid
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336
Bacillus pumilus
HO H3CO
HO H3CO
COOH ferrulic acid
vinylguaiacol
Scheme 48.
Baicalin (baicalein-7-O-glucuronide) and baicalein (5,6,7-trihydroxyflavone) are the most important biologically active components of Radix Scutellariae. They exhibit wide range of pharmacological activities due to their antiallergic, anti-inflammatory, antiatherogenic, antithrombotic, antibacterial, antiviral and anticancerogenic properties. These properties arise mainly from antioxidant activity of flavonoids and may be changed or improved by modification of compound structures. Several microorganisms not only hydroxylate balcalein but also catalyze the reaction of its O-methylation (Scheme 49) yielding various flavonoids of modified biological properties [90]. Kinetic resolution is a well-established method for the preparation of optically active compounds and was described widely in this review. However, it has a disadvantage in that the maximum yield of the desired enantiomer is theoretically limited to 50%. Also the separation of the product and the recovered starting material is inevitable, which sometimes is tedious. Racemization offers the mean of preparation of enantiomerically pure products in 100% yield. As an example may serve racemization of 2-arylpropanoic acids by Nocardia diaphanozonaria, a reaction which was also used to prepare enantiomerically pure (R)ibuprofen, a non-steroidal anti-inflammatory drug (Scheme 50), in excellent yield [91]. OCH3 HO
O
HO
+
H3CO
H3CO
OH O
OH O
Penicilium chrysogenum
HO
O Chaetomium sp.
HO
O
HO H3CO
OH O
OH O
baicalein Coryneum betulinum
OH HO
O
HO OH O
Scheme 49.
O
Application of Whole-cell Biocatalysis in Chemoenzymatic…
337
CH3
CH3 COOH
Nocardia diaphanozonaria
COOH
Scheme 50.
Synthesis of L-carnitine is an another, excellent example of the application of dercemizations catalyzed by microbial cells. This compound is essential for the fatty acid transportation through the mitochondrion plasma membrane and therefore influences the lipids metabolic pathways and, going further, is applied as a diet supplement in heart diseases or in a carnitine deficiency syndrome. Chemical synthesis of carnitine results in racemic mixture of this compound, which is then resolved via chemical derivatization [92]. Enantiomeric D-carnitine is a waste-product of this process. This enantiomer is converted in cells of many bacteria, such as Escherichia, Salmonella or Proteus into the opposite Lisomer, in quite complicated manner (Scheme 51), by intracellular enzymes, which are induced only under anaerobic conditions. Unfortunately under such conditions the valuable L- carnitine is subsequently metabolized inside the living cells. The mentioned problem was overcame by cells pre-treatment, by simple addition of some amount of L-carnitine to the cultivation medium, which cause anaerobic enzyme induction in the cells of Escherichia coli. Then the bioconversion of (D)-enantiomer into desired L-carnitine was carried out aerobically [93] (Scheme 22). Quite unusual is racemization of benzoin catalyzed by Rhizopus oryzae (Scheme 52). When reaction was carried out in the range of pH of 4-5 S-isomer was obtained with high yield. Changing pH to 7.5-8 cause complete reveal of enantiospecificity and enantiomerically pure R-isomer is produced. OH D-carnitine
N
+
COO-
D-carnitine dehydrogenase
racemase
NADH NAD OH
L-carnitine
Scheme 51.
N+
COO-
O N
+
COO-
dehydrokarnityna L-carnitine dehydrogenase
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338
O
OH Rhizopus oryzae pH 7.5-8
Rhizopus oryzae pH 4-5
O
O
OH
Rhizopus oryzae pH 7.5-8
OH
Scheme 52.
5. References [1] [2] [3] [4] [5] [6] [7] [8] [9]
[10] [11] [12]
[13]
A.M. Roughi, Chiral business, Chem. Eng. News (2003) 81, 45-55. H. Caner, E. Groner, L. Levy, I. Agranat, Trends in the development of chiral drugs, Drug Discovery Today (2004) 9, 105-110. A. Schmid, J. S. Dordick, B. Hauer, A. Kiener, M. Wubbolts, B. Witholt, Industrial biocatalysis today and tomorrow, Nature (2001) 409, 258-268. B. G. Davis, V. Boyer, Biocatalysis and enzymes in organic synthesis, Nat. Prod.Rep. (2001) 18, 618-640. T. P. Begley, M.-D. Tsai, Biocatalysis and biotransformation. Enzymology in the genomic era. Curr. Opin. Chem. Biol., (2003) 7, 228-229. A. M. Roughi, Chem. Eng. News (2001) 79, 37-43. D. J. Pollard, J. M. Woodley, Biocatalysis for pharmaceutical intermediates; the future is now, Trends Biotechnol (2006) 25, 66-73. H. G. W. Leuenberger, Biotransformation – a useful tool in organic chemistry, Pure & Appl. Chem. (1990) 62, 753-768. R. de Souza Pereira, The use of the baker’s yeast in the generation of asymmetric centers to produce chiral drugs and others compounds. Crit. Rev.Biotechnol. (1998) 1, 25-83. O. Ward, C. Young, Reductive biotransformations of organic compounds by cells or enzyme of yeast. Enzyme Microb. Technol. (1990) 12, 482-503. R. Csuk, B. Glanzer, In Stereoselective Biocatalysis, R.N. Patel Edition Marcel and Dekker, Inc. New York (2000), 527-578. R. Patel, A. Banarejee, C. McName, D. Brzozowski, R. Hanson, L. Szarka, Highly Regio- and Enantioselective Reduction of 3,5-Dioxocarboxylates. Enzyme Microb. Technol. (1993) 15, 1014-1021. R.F. Brown, W.R. Jackson, T.D. McCarthy, A synthesis of (-)-denopamine. Tetrahedron: Asymm. (1993) 4, 2149-2150.
Application of Whole-cell Biocatalysis in Chemoenzymatic…
339
[14] J. Goswami, R.L. Bezbaruah, A. Goswami, N. Borthakur, A convenient steroselective synthesis of (R)-(-)-denopaine and (R)-(-)-salmeterol. Tetrahedron: Asymm. (2001) 12, 3343-3348. [15] J.A. Pesti, R. DiCosimo, Recent progress in enzymatic resolution and desymmetrization of pharmaceuticals and their intermediates. Curr. Opin. Drug Discov. Develop. (2003) 6, 884-901. [16] M.P Burns, J.W. Wong, Microbial asymmetric reduction of 2-chloro-1-[-6-(2,5dimethyl-pyrrol-1-yl)-pyridin-3-yl]-ethanone. (2002) Patent US-06451587, Pfizer Inc. [17] L. Banfi, G. Cascio, C. Ghiron, G. Guanti, E. Manghisi, E. Narisano, R. Riva, Microbiological enantioselective synthesis of (S) and (R) 4-(p-anisyloxy)-3hydroxybutyrates as a new chiral building blocks for the synthesis of β-lactam antibiotics. Tetrahedron: Asymm. (1994) 50, 11983-11994. [18] R. Patel, L. Chu, R. Chidambaran, J. Zhu, J. Kant, Enantioselective microbial reduction of 2-oxo-2-(1′,2′,3′,4′-tetrahydro-1′,1′,4′,4′-tetramethyl-6′naphthalenyl)acetic acid and its ethyl ester. Tetrahedron: Asymm. (2002) 12, 349-355. [19] G. Fronza, C. Fuganti, P. Graselli, A. Mele, On the mode of baker’s yeast transformation of 3-chloropropiophenone and related ketones . Synthesis of (2s)-[22H]propiophenone, (R)-fluoxetine, and (R) and (S) fenfluramine. J. Org. Chem. (1991) 56, 6019-1023. [20] A. Kumar, D.H. Ner, S. Dike, A novel chemoenzymatic enantioselective synthesis of some clinically effective CNS drugs and related compounds. Indian J. Chem (1992) 31B, 803-809. [21] T.M. Koening, D. Mitchel, A convenient method for preparing enantiomerically pure norfluoxetine, fluoxetine and tomexetine. Tetrahedron Lett. (1994) 35, 1339-1342. [22] R.J. Hammond, B.W. Poston, I. Ghiviriga, B.D. Feske, Biocatalytic synthesis towards both antipodes of 3-hydroxy-3-phenylpropanitrile a precursor of fluxetine, amoxetine and nisoxetine. Tetrahedron Lett. (2007) 48, 1217-1219. [23] R.N. Patel, L. Chu, R. Mueller, Diastereoselective microbial reduction of (S)-[3chloro-2-oxo-1-(phenlmethyl)propyl]carbamic acid, 1,1-dimethyl ester. Tetrahedron: Asymm. (2003) 14, 3105-3109. [24] G.L Drusano, J.A. Libello, S.L. Preston, E. O’Mara, S. Kaul, S. Schnittman, R. Echols, Hollow-fiber unit evaluation of a new human immunodeficiency virus type 1 protease inhibitor, BMS-232632, for determination of the linked pharmacodynamic variable. J. Infect. Diseas. (2001) 183, 1126-1129. [25] R.M. Patel, Bicatalysis: Synthesis of Chiral intermediates for pharmaceuticals. Current Organic Chemistry (2006) 10, 1289-1321. [26] R. Patel, A. Banerjee, C. McName, D. Brzozowski, L. Szarka, Preparation of chiral synthon for HIV protease inhibitor: stereselective microbial reduction of N-protected -aminochloroketone. Tetrahedron: Asymm. (1997) 8, 2547-2552. [27] H. Matsumae, H. Douno, S. Yamada, T. Nishida, Y. Ozaki, T. Shibatani, T. Tosa, Microbial asymmetric reduction of (RS)-2-(4-methoxyphenyl)-1,5-benzothiazepin3,4(2H,5H)-dione. J. Ferment. Bioeng (1995) 79, 28-32. [28] R. DiCisimo, P.Ma, J.A. Pesti, Enantioselective enzymatic aminolysis of a racemic 2isoxazolylacetatate alkyl ester. (2002) Patent US-2002160465, Bristol-Myers Squibb.
340
Ewa Żymańczyk-Duda and Paweł Kafarski
[29] W. Adam, Z. Lucacs, C. R. Saha-Möller, P. Schreier, Biocatalytic kinetic resolution of racemic hydroperoxides though the enantioselective reduction with free and immobilized microorganisms, J. Am Chem. Soc. (2000) 122, 4887-4892. [30] D. Graham, R. Holt, P. Lindberg, S. Tylor, Enantioselective preparation of pharmaceutically active sulfoxides by bioreduxtion. (2003) Patent EP-00795025, AstraZeneca AB. [31] W. Kromer, The relationship between pH-selectivity, chirality and efficacy of gastric proton pomp inhibitors. Curr. Top. Pharmacol. (2000) 5, 45-69. [32] A.Zaks, Industrial Biocatalysis. Current Opinion in Chemical Biology (2001) 5, 130136. [33] R.L. Hanson, J.M. Howell, T.L. LaPorte, M.J. Donavan, D.L. Cazzulino, V. Zonella, M.A. Montana, V.B. Nanduri, S.R. Schwartz, R.F. Eiring, Synthesis of allysine ethylene acetal using phenylalanine dehydrogenase from Thermoactinomycetes intermedius. Enzyme Microb. Technol. (2000) 26, 348-358. [34] R. N. Patel, Enzymatic synthesis of intermediates for Omapatrilat, an antihypertensive drug, Biomolec. Eng. (2001) 17, 167-182. [35] P. Prave, U. Faust, W. Sitting, D.A. Sukatsch, Fundamentals of Biotechnology. VCH Publishers, Weinhem 563-600. [36] M. Miyauchi, R. Endo, M. Hisaoka, H. Yasuda, I. Kawamoto, Synthesis and structureactivity relationship of a novel oral carbapenem, CS-834. J Antibiot (Tokyo) (1997) 50, 429-439. [37] J.F. Almeida, J.Anaya, N. Martin, M. Grande, J.R. Moran, M.C. Caballero, New enantioselective synthesis of 4-hydroxy-2-oxopyrrolidine-N-acetamide (oxiracetam) from malic acid. Tetrahedron: Asymm. (1992) 3, 1421-1430. [38] J. Multzer, R. Zuhse, R. Schmiechen, Enantioselective synthesis of the antidepressant rolipram via Michael addition to a nitro olefin. Angew. Chem. Int. Ed (1992) 31, 870872. [39] M. Orena, G. Porzi, S. Sandri, A New synthesis of (R)-GABOB and (R)-oxiracetam. J. Chem. Res. (Synopses) (1990) 11, 376-378. [40] J.Aube, Y. Wang, S. Ghosh, K.L. Langhaus, Oxaziridine-mediated ring expansions of substituted cyclobutanones: Synthesis of (-)-γamino-β-hydroxybutyric acid (GABOB). Synth. Commun. (1991) 21, 693-701. [41] B. Buckland, D. Robinson, M. Chrtrain, Biocatalysis for pharmaceuticals – status and prospects for a key technology. Metab. Eng. (2000) 2, 42-48. [42] H. Berner, H. Vyplel, G. Schultz, P. Struchlik, Chemistry of pleuromutilins, IV. Synthesis of 14-O-acetyl-8-alpha-hydroxy-mutilins. Tetrahedron (1983) 39, 13171321. [43] D. Żelaszczyk, K. Kieć-Konowicz, Biocatalytic approachesβ-Blockers. Curr. Med. Chem. (2007) 14, 53-65. [44] S.L Jonstone, G.T. Philips, B.W. Robertson, P.D. Watts, M.A. Bertola, H.S. Koger, A.F. Max. In Biocatalysis in Organic Media, C. Laane, J. Tramper, M.D. Lilly Edition, Elsevier Amsterdam (1987), 387-392. [45] R. Patel, A. Banerjee, C. McNama, L. Szarka, Stereoselective epoxidation of 2,2dimethyl-2H-1- benzopyran-6-carbonitrile. J. Bioorg. Med. Chem. (1994) 2, 535-542.
Application of Whole-cell Biocatalysis in Chemoenzymatic…
341
[46] J. Das, M. Haslanger, J. Gougoutas, M. Mallley, Synthesis of Optically Active 7Oxabicyclo[2.2.1]heptanes and Assignment of Absolute Configuration. Synthesis (1987) 12, 1100-1112. [47] P. Villeneuve, J. M. Muderhwa, J. Graille, M. J. Haas, Customizing lipases for biocatalysis: a survey of chemical, physical and molecular biological approaches. J. Mol. Catal. B (2000) 9, 113- 1448. [48] G. Grogan, Biotransformations, Annu. Rep. Prog. Chem. B (2005) 101, 192-209. [49] K. E. Jaeger, B. W Dijkstra, M. T. Reetz, Bacterial biocatalysts: molecular biology, three-dimensional structures and biotechnological application of lipases. Annu. Rev. Microbiol. (1999) 53, 315-351. [50] G. Fantin, M. Fogagnolo, A. Guerrini, A. Medici, P. Pedrini, S. Fontana, Enantioselectyive hydrolyses with Yarrowia lipolytica: A versatile strain for esters, enol esters, epoxides, and lactones. Tetrahedron: Asymm. (2001) 12, 2709-2713. [51] V. Gotor-Fernández, E. Busto, V. Gotor, Candida antarctica lipase B: an ideal biocatalyst for preparation ofnitrogenated organic compounds. Adv. Synth. Catal. (2006) 348, 797-812. [52] S. Tamalampudi, S. Hama, T. Tanino, M. L. Talukder, A. Kondo, H. Fukuda, Development of recombinant Aspergillus oryzae whole-cell biocatalyst expressing lipase-encoding gene from Candida Antarctica. Appl. Microbiol. Biotechnol. (2007) 75, 387-395) [53] S. Tamalampudi, S. Hama, T. Tanino, M. L. Talukder, A. Kondo, H. Fukuda, Immobilized recombinant Aspergillus oryzae expressing heterologous lipase: An efficient whole-cell biocatalyst for enatioselective transesterification in non-aqueous medium. J. Mol. Catal. B (2007) 48, 33-37. [54] A. S. Demir, H. Hamamci, C. Tanyeli, I. M. Akhmedov, F. Doganel, Synthesis and Rhizopus oryzae mediated enenatioselective hydrolysis of -acetoxy aryl alkyl ketones. Tetrahedron: Asymm. (1998) 9, 1673-1677. [55] U. R. Kalkote, S. R. Ghorpade, R. R. Joshi, T. Ravindranathan, K. B. Bastawade, D. V. Gokhale, A practical and scalable process for 4-(R)-cyclopent-2-en-1-(S)-acetate by dyssymetrization of meso-cyclopent-2-en-1,4-diacetate catalyzed by Trichosporon beigelii (NCIM3326), a cheap biocatalysis. Tetrahedron: Asymm. (2000) 11, 29652970. [56] A. Romano, D. Romano, F. Molinari, D. Gandolfi, F. Constantino, A new chemnoenzymatic synthesis of D-cloprostenol, Tetrahedron: Asymm. (2005) 16, 32793282. [57] D. Romano, F. Falconi, D. Mora, F. Molinari, A. Buthe, M. Ansorge-Schumacher, Enhanced enetioselectivity of Bacillus coagulans in the hydrolysis of 1,2-Oisopropylideneglycerolesters by thermal knock-out of undesired enzymes. Tetrahedron: Asymm. (2005) 16, 841-845. [58] A. Miyadera, A. Imura, Enantioselective synthesis of a key intermediate of Levofloxacin using microbial resolution. Tetrahedron: Asymm. (1999) 10, 119-123. [59] K. Laumen, M. Kittelmann, O. Ghisalba, Chemo-enzymatic approaches fort he creation of novel chiral building blocks and reagents for pharmaceutical applications. J. Mol. Catal. B (2002) 19/20, 55-66.
342
Ewa Żymańczyk-Duda and Paweł Kafarski
[60] L. Martinková, N. Klempier, I. Prepechalová, V. Prikrylová, M. Ovsená, H. Griengl, V, Kren, Chemoselective biotransformation of nitriles by Rhodococcus equi A4. Biotechnol. Lett. (1998) 909-912. [61] W. Kroutil, I. Osprian, M. Mischitz, K. Faber, Chemoenzymatic synthesis of (S)-(-)Frontalin using bacterial epoxide hydrolases. Synthesis (1997) 156-158. [62] S. J. C. Taylor, A. G. Sutherland, C. Lee, R. Wisdom, S. Thomas, S. M. Roberts, C. Evans, Chemoenzymatic synthesis of (–)-carbovir utilizing a whole cell catalysed resolution of 2-azabicyclo[2.2.1]hept-5-en-3-one. Chem. Commun. (1990) 1120-1121. [63] K. Sakamo, K. Honda, K. Wada, S. Kita, K. Tsuzaki, H. Nose, M. Kataoka, S. Shimizu, Practical resolution system for D,L- pantoyl lactone using lactonase from Fusarium oxysporium. J. Biotechnol. (2005) 118, 99-106. [64] S. R. Wallner, B. M. Nestl, K. Faber, Highly enantioselective sec-alkyl sulfatase activity of Sulfolobus acidocaldarius DSM 639. Org. Lett. (2004) 5009-5010. [65] C. Mateo, B. Fernandes, F. van Rantwijk, A. Stolz, R. A Sheldon, Stabilization of oxygen-labile nitrilases via co-aggregation with poly(ethyleneimine). J. Mol. Catal. B (2006) 38, 154-157. [66] O. Meth-Cohn, M.-X. Wang, An in depth study of the biotransformation of nitriles imto amides and/or acids using Rhodococcus rhodochrous AJ270. J. Chem. Soc. Perkin1 Trans. (1997) 1099-1104. [67] M.-X. Wang, G.-Q. Feng, Enzymatic synthesis of optically active 2-m3thyl- and 2,2dimethylcyclopropanecarboxylic acids and their derivatives. J. Mol. Catal. B (2002) 18, 267-272. [68] M.-X. Wang, G.-Q. Feng, A novel approach to enentiopure cyclopropane compounds from biotransformation of nitriles. New. J. Chem. (2002) 26, 1575-1583. [69] Y.-F. Tang, J.-H. Xu,Q. Ye, B. Schulze, Biocatalytic hydrolysis of (S)-phenyl glycidyl ether using newly isolated Baciullus megaterium ECU1001. J. Mol. Catal. B (2001) 13, 61-68. [70] S. Pedragosa-Moreau, R. Archelas, R. Furstoss, Microbiological transformations. 28. Enantioceomplememtary epoxide hydrolases as a preparative access to both enantiomers of styrene oxide. J. Org. Chem. (1993) 58, 5533-5536. [71] J.-P. Chao, H. Fu, T.-E. Lo,P. T. Chen, J.-J. Wang, One-step production of D-phydroxyphenylglycine by recombinant Escherichia coli strains, Biotechnol. Prog. (1999) 15, 2039-2045. [72] S. Martinez-Rodriguez, F. J. Las Heraz-Vasquez, J. M. Clemente-Jimenez, L. Mingorance-Cazarola, F. Rodriguez-Vico, Complete conversion of D,L-5monosubstitutedhydantoins with a low velocity of chemical racemization into D-amino acids using whole cells of recombinant Escherichia coli. Biotechnol. Prog. (2002) 18, 1201-1206. [73] X. Chen, Y. Zheng, Y. Shen, A new method for production of valienamine with microbial degradation of acarbose, Biotechnol. Prog. (2005) 21, 1002-1003. [74] W. D. Fessner, Enzyme-mediated C-C bond formation, Curr. Opinion Chem. Biol. (1998) 2, 85-97. [75] V. K. Krieble, Enzymes. Asymmetric syntheses through the action of oxynitrilases, J. Am. Chem. Soc. (1913) 35, 1643-1747.
Application of Whole-cell Biocatalysis in Chemoenzymatic…
343
[76] H. Griengl, H. Schwab, M. Fechter, The synthesis of chiral cyanohydrins by oxynitrilases, Trends Biotechnol. (2000) 18, 252-256. [77] L. Synoradzki, T. Rowicki, M. Włostowski, Calcium pantothenate. Optimisation of oxynitrilase-catalysed asymmetric hydrocyanation of 3-hydroxy-2,2-dimethylaldehyde: synthesis of (R)-pantolactone. Org. Proc. Res. Dev. (2006) 10, 103-108. [78] F. Effenberger, J. Jäger, Synthesis of adrenergic bronchodilators (R)-terbutaline and (R)-salbutamol from (R)-cyanohydrins. J. Org. Chem. (1997) 62, 3867-3873. [79] G. Ritčenkovs, L. T. Kanerva, Otimized methods for the preparations of (S)-2hydroxy-2-phenylpropanenitrile exploiting (R)-oxynitrillase in almond meal. L. Mol. Catal. B (2000) 11, 37-43. [80] A. Solis, H. Luna, H. I. Perez, N. Manjarrez, Evaluation of guanabana (Annona muricata) seed meal as a source of (S)-oxynitrilase. Tetrahedron: Asymm. (2003) 14, 2351-2353. [81] S.-Q. Han, P.-K. Ouyang, P. Wei, Y. Hu, Enzymatic synthesis of (R)-cyanohydrins by a novel (R)-oxynitrilase from Vicia sativa L.. Biotechnol. Lett. (2006) 28,1909-912. [82] H. J. M. Gijsen, L. Qiao, W. Fitz, C.-H. Wong, Recent advances in the chemoenzymatic synthesis of carbohydrates and carbohydrate mimetics. Chem. Rev. (1996) 96, 443-473. [83] G. Feron, G. Mauvais, F. Martin, E. Sémon, C. Blin-Perrin, Microbial production of 4hydroxybenzilidene acetone, the direct precursor of raspberry ketone. Lett. Appl. Microbiol. (2007) 45, 29-35 [84] T. H. Kurniadi, R. B. Rhlid, M. A. Juillerad, M. Schüler, R. G. Berger, Enantiogenic synthesis of (R)-(-)-3-hydroxy-1-penten-4-one. Tetrahedron: Asymm. (2003) 14, 363366. [85] M. Pohl, B. Lingen, M. Müller, Thiamin-diphosphate-dependent enzymes: new aspects of asymmetric C-C bond formation. Chem.-Eur. J. (2002) 23, 5288-5295. [86] T. Matsuda, Y.Ohashi, T. Harada, R. Yanagifara, T. Nagasawa, K. Nakamura, Conversion of pyrrole to pyrrole-2carboxylate by cells of Bacillus megaterium in supercritixcal CO2. Chem. Commun. (2001) 2194-2195 [87] M. Aresta, A. Denebedetto, Dalton Trans. (2007) 90, 2975-2992. [88] R. H. Cichewicz, S. A. Kouzi, Trasformation of resveratrol to piceid by Bacillus cereus. J. Nat. Prod. (1998) 61, 1313-1314. [89] I.-Y. Lee, T. G. Volm, J. P. N. Rosazza, Decarboxylation of ferrulic acid to 4vinylguaiacol by Bacillus pumilus in aqueous-organic solvent two-phase system. Enzyme. Microb. Technol. (1998) 23, 261-266. [90] E. Kostrzewa-Susłow, J. Dmochowska-Gładysz, J. Oszmiański, Microbial transformation of baicalin and baicalein. J. Mol. Caral. B (2007) 49, 113-117. [91] D.-I. Kato, S. Mitsuda, H. Ohta, Microbial deracemization of -substituted cvarboxylic acids: substrate specificity and mechanistic investigation. J. Org. Chem. (2003) 68, 7234-7242. [92] J.M. Obon, J.M. Maiquez, M. Canovas, H.P. Kleber, J.L. Iborra, L(-)-carnitine productionwith immobilized Escherichia coli cells in continuous reactors. Enzyme Microb. Technol. (1997) 21, 531-536.
344
Ewa Żymańczyk-Duda and Paweł Kafarski
[93] M.R. Castellar, M.Canovas, H.P. Kleber, J.L. Iborra, Biotransformation of D-(+)carnitine into L(-)-carnitine by resting cells of Escherichia coli O44 K74. J. Appl. Microb. (1998) 85, 883-890.
Index
1 1-butyl-3-methylimidazolium, xiii, 245, 252
3 3,4-ethylenedioxythiophene (EDOT), xiii, 245, 247, 248, 249, 250, 255, 257, 258
A abiotic, 266 absorption spectra, 26 acarbose, 342 access, 22, 104, 160, 185, 201, 213, 260, 331, 342 accounting, 102, 297 accuracy, 23, 28 acetic acid, 181, 313, 339 acetone, 105, 332, 333, 343 acetonitrile, 81, 134, 172 acetylcholinesterase, 23 acidic, 62, 87, 88, 111, 145, 198, 249, 253, 270 acidification, 284 acrylic acid, 29 activated carbon, 290, 291, 299 activated sludge flocs, 129, 131 activation, vii, 1, 49, 80, 81, 109, 159 active site, 3, 28, 54, 55, 56, 58, 59, 60, 62, 63, 81, 107, 160, 162, 180, 188, 189, 191, 201, 329 activity level, 131 actuation, 121 acylation, 75, 175 adaptation, 211, 260, 261, 262, 271
additives, 172, 196, 265 adenine, 54, 68, 72, 74 adenosine, viii, 48, 64, 72, 73, 78, 84, 86, 87, 120 adenosine deaminase (ADA), 64, 71, 72, 73 adenosine triphosphate (ATP), 85, 86, 87, 120 adhesion, 285 adsorption, vii, 1, 3, 17, 20, 21, 24, 26, 27, 28, 29, 31, 33, 52, 68, 69, 70, 81, 89, 100, 121, 130, 131, 270, 272, 279, 283, 284, 290, 291 aerobe, 271 aerobic bacteria, 116 aerospace, 3 Africa, 117 agar, 113, 114, 115, 284 Agaricus bisporus, 106, 134, 143 age, 107, 167 agent, 49, 75, 101, 107, 145, 196, 200, 225, 278, 284, 315, 319, 320, 324, 328 aggregates, vii, 1, 3, 246 aggregation, 246, 253, 342 aging, 107 agonist, 310, 311, 313 agriculture, 223 agroindustrial, ix, 95, 138 AIDS, 323 alanine, 196 albumin, 111, 278 alcohol(s), 24, 76, 78, 132, 138, 148, 166, 170, 172, 177, 184, 196, 205, 243, 260, 309, 310, 312, 313, 322, 327 alcoholism, 313 alcoholysis, ix, 76, 77, 78, 155, 160, 161, 170, 172, 181 aldehydes, 331 aldolase, 333 algae, 176, 260, 262
346
Index
algorithm, 193, 266, 277 alkaline, xi, 24, 87, 88, 110, 122, 142, 144, 145, 146, 150, 152, 209, 210, 211, 212, 214, 215, 216, 217, 218, 219, 220, 221, 222, 223, 224, 225, 226, 227, 228, 265, 273, 276, 280 alkaline phosphatase, 24, 88, 273, 280 alkalinity, xi, 209 alkaliphiles, xi, 219, 221, 224, 225, 226 alkaliphilic actinomycetes, xi, 219, 220, 221, 223, 224, 225 alkylation, 313 allergens, 151 allylamine, 29 alternative(s), viii, ix, xiv, xv, 47, 48, 53, 63, 72, 73, 95, 96, 98, 111, 112, 113, 124, 129, 133, 134, 136, 149, 152, 159, 166, 173, 176, 184, 187, 200, 246, 247, 251, 273, 308, 313, 329 alters, 180 amide, 325, 329 amines, 107, 131, 132, 260, 322, 324, 325 amino, 54, 62, 66, 69, 78, 88, 189, 212, 224, 260, 263, 265, 271, 273, 310, 316, 319, 329, 330 amino acid(s), 54, 62, 88, 189, 212, 224, 260, 263, 265, 271, 329, 330 ammonia, 35, 289, 327 ammonium, 87, 104, 226, 270, 334 ammonium sulphate, 104 amylase, xi, 122, 142, 148, 153, 209, 211, 212, 215, 216, 219, 223, 225, 264 anaerobic bacteria, 118 anaerobic digesters, 143 anaerobic sludge, 119, 120, 124, 125, 143, 147, 148, 285, 286 angina, 320 angiogenesis, 278 aniline, 246, 247, 248, 251, 252, 253, 254, 255, 290, 291 animal models, 156 animals, 100, 102, 103, 105, 113, 158, 160, 162, 223 anion, 68, 87, 278 antagonists, 311 anthracene, 140 antibacterial, 320, 336 antibiotic(s), xi, 97, 209, 210, 218, 219, 222, 226, 239, 243, 270, 274, 284, 285, 312, 319, 324, 328, 339 antibiotic resistance, xi, 209, 210 antibody, 192, 278 anti-cancer, 270 anticancer drug, 222, 313
anticoagulant, 273 anticonvulsant, 319 antidepressant, 313, 319, 328, 340 anti-inflammatory, 156, 336 antimetabolites, viii, 47 antioxidant(s), 29, 146, 180, 336 antitumor, 49, 222, 226, 270 antiviral, viii, 47, 48, 72, 85, 336 antiviral agents, 85 antiviral drugs, viii, 47 anxiety, 156, 313 apoptosis, 149 aqueous solutions, 24, 106, 107, 147, 252 arachidonic acid, 176, 183 archaea, xi, 209, 211, 212, 213, 217, 218, 220, 221, 264, 276 arginine, 35, 58, 60, 113, 244 argon, 13 aromatic compounds, 109, 335 aromatic hydrocarbons, 214, 291 arteriosclerosis, 315 Asia, 117 Aspergillus niger, 104, 116, 117, 137, 144, 145, 147, 148, 150, 166, 315, 329 assessment, xi, 210, 239, 243 assignment, 227 assimilation, 102, 127, 128 assumptions, 21, 288 asthma, 332 asymmetric synthesis, 322 asymmetry, xiv, 308 atmospheric pressure, 161, 334 atomic force microscopy (AFM), xiii, 28, 35, 245, 248, 250 atoms, 11, 63, 109, 120, 318 attachment, 3, 11, 16, 19, 275, 283, 284, 285, 287, 290 attacks, 162 attention, xi, 6, 22, 51, 97, 115, 131, 136, 168, 209, 210, 215, 219, 221, 222, 246, 249, 273, 283 availability, 51, 52, 100, 101, 120, 164, 261, 266, 273 awareness, 211 azo dye, 131, 144
B bacillus, 54, 68, 105, 113, 116, 118, 122, 145, 148, 150, 152, 191, 212, 216, 217, 218, 226, 227, 228, 263, 275, 295, 315, 324, 329, 334, 335, 341, 343
Index bacillus subtilis, 54, 68, 122, 191, 212, 315 bacteria, xi, xii, 85, 96, 103, 104, 105, 106, 108, 114, 116, 118, 120, 122, 141, 142, 160, 209, 210, 211, 214, 215, 216, 217, 220, 222, 223, 224, 226, 230, 231, 260, 261, 262, 263, 265, 271, 274, 275, 277, 280, 284, 285, 287, 315, 333, 334, 337 bacterial cells, 87, 284, 285 bacterial fermentation, 113 bacteriolysis, 243 bacterium, 113, 122, 140, 149, 153, 215, 216, 217, 270, 271, 276, 278, 279 baicalein, 336, 343 barriers, 14, 16 base pair, 212 basidiomycetes, 320 baths, 132, 146 batteries, 246 beef, 324 behavior, 10, 130, 131, 140, 165, 172, 217, 289, 291, 294, 299 beneficial effect, 171 benefits, ix, xii, 84, 113, 156, 162, 164, 168, 229 benign, 164, 188, 204, 246 benzene, 214, 246, 253 benzodiazepine, 181 bicarbonate, 87 binding, 34, 55, 56, 57, 58, 59, 60, 62, 113, 162, 180, 270, 272, 283, 284 bioactive compounds, xi, 219, 260 bioavailability, 103, 120, 261 biocatalyst(s), viii, x, ix, xi, xii, xiii, xiv, xv, 2, 3, 28, 31, 36, 48, 51, 52, 53, 63, 64, 66, 67, 68, 69, 80, 81, 87, 95, 96, 98, 100, 106, 112, 133, 134, 155, 167, 168, 169, 164, 187, 189, 190, 192, 196, 197, 199, 200, 201, 202, 204, 207, 210, 211, 215, 219, 222, 225, 229, 235, 241, 243, 246, 259, 260, 263, 265, 281, 282, 283, 284, 287, 297, 298, 308, 309, 316, 318, 320, 322, 333, 341 biocatalytic process, vii, xiv, 1, 2, 35, 165, 251, 308, 321, 327 bioconversion, ix, 70, 155, 191, 241, 309, 313, 314, 337 bioconversions, xv, 51, 52, 68, 89, 308 biodegradability, viii, 95, 96, 119, 128, 148, 152 biodegradable, 119, 130, 131, 133, 291 biodegradation, 113, 119, 120, 121, 130, 136, 138, 145, 148, 150, 218, 285, 286, 289, 290, 291 biodiesel, 2, 110, 140, 144 biodiversity, 101, 213, 225, 260, 277
347
biofilms, 213, 262, 284, 285, 286, 287, 289, 299, 300, 302 biofuel(s), vii, 2 biological activity, 273 biological nutrient removal, 141 biological processes, 51, 220 biological systems, 288 biological toxicity, 214 bioluminescence, 275 biomass, 113, 116, 119, 120, 121, 123, 124, 125, 127, 139, 151, 178, 223, 240, 242, 267, 269, 270, 271, 282, 285, 286, 287, 290 biomass growth, 119 biomolecules, 8, 25, 26, 28, 29, 220, 221, 225, 277 biopolymer(s), 24, 107 bioremediation, 2, 106, 188, 235 biosensors, 6, 22, 109, 267 biosphere, 260, 277 biosynthesis, 148, 176, 206, 243, 277 biosynthetic pathways, 53 biotechnology, 3, 89, 143, 145, 147, 192, 214, 215, 218, 220, 221, 225, 226, 231, 267, 276, 277, 299, 305 biotransformations, xv, 53, 79, 80, 184, 260, 308, 309, 325, 328, 338 Black Sea, 271, 278 bleaching, 132, 140, 149 blocks, 331 blood, 173, 273, 280, 309 blood group, 273, 280 body temperature, 171 bonding, 60, 63 bonds, 10, 60, 68, 115, 162, 163, 169, 266, 283, 333 bottom-up, vii, 1, 3, 4 bounds, 100 brain, 156 Braun’s lipoprotein, xii, 230, 231, 233, 239 Brazil, 95, 116, 117, 138, 139, 150 breakdown, 84, 117 bromination, 266 bromine, 266 bronchitis, 324 bronchospasm, 332 buffer, 26, 28, 32, 36, 71, 76, 77, 79, 81, 135, 166, 202, 246, 272, 334 building blocks, 3, 325, 328, 329, 339, 341 Bulgaria, 281 Bupropion, 322 Burkholderia, 173 by-products, viii, 47, 48, 50, 103, 115, 266, 327
Index
348
C Ca2+, 103 cachexia, 156, 175 caffeine, 115, 116, 136, 139, 149 calcium, x, 21, 29, 144, 195, 196, 197, 199, 200, 203, 204, 205, 284, 315, 326 calcium carbonate, 29 calcium channel blocker, 315 calcium pantothenate, x caloric intake, 160 calorie, 169, 178 Canada, 173 cancer, 222, 278, 328 cancer cells, 222 candidates, xiv, 115, 136, 211, 212, 307 capillary, 17 capsule, 32, 33, 35 carbohydrate(s), viii, 48, 49, 53, 115, 118, 170, 139, 343 carbon, 53, 65, 97, 100, 101, 102, 103, 113, 120, 131, 135, 141, 142, 156, 158, 163, 164, 166, 167, 170, 171, 172, 173, 174, 181, 185, 214, 257, 261, 266, 290, 291, 331, 333, 334, 335 carbon atoms, 120, 158 carbon dioxide (CO2), 123, 164, 166, 167, 172, 173, 174, 181, 184, 185, 203, 334, 335, 343 carbonyl groups, 60 carboxylic acids, 327 carcinoma, 279 cardiac arrhythmia, 320 cardiovascular disease, 156, 175 carrageenan, 199, 200, 205 carrier, 203, 270, 283, 284, 286, 287, 289, 295 case study, 289 casein, 215 casting, 252 castor oil, 111, 140, 173, 185 catalase, 23, 29, 31, 205, 262 catalysis, xiv, 35, 36, 58, 62, 63, 91, 162, 166, 171, 180, 181, 188, 189, 205, 244, 256, 275, 308 catalyst(s), vii, ix, xii, xiii, xiv, xv, 2, 51, 52, 54, 70, 77, 80, 81, 85, 86, 89, 134, 144, 146, 148, 155, 159, 166, 172, 173, 189, 197, 200, 203, 206, 211, 229, 237, 242, 246, 250, 256, 259, 281, 292, 296, 308, 321, 323, 329, 333 catalytic activity, 3, 28, 29, 31, 32, 34, 35, 36, 52, 66, 88, 263, 275, 292 catalytic hydrogenation, 171 catalytic properties, 2, 52, 152
catalytic system, xiii, 245, 247, 251 cation, 109, 138, 224, 253, 272 cattle, 324 cDNA, 206, 214, 267 cell adhesion, 285 cell culture, 278, 284, 292 cell death, 294 cell division, 285 cell growth, xii, xiv, 230, 241, 262, 281, 285, 293, 294, 297, 299 cell membranes, 235, 237, 238, 284 cell metabolism, 293, 294 cell surface, 111, 283 cell wall, x, 196, 269, 283 cellulose, 29, 30, 35, 87, 116, 122, 223, 284 central nervous system (CNS), 175, 339 ceramic(s), 24, 283 cereals, 98 cervical cancer, 149 channels, 231, 285 chemical composition, 152 chemical energy, 160 chemical industry, 132, 159, 170, 177 chemical reactions, ix, 2, 187, 320 chemical reagent, x, 196, 201 chemical stability, 68 chemoenzymatic processes, xv, 308 chemotherapy, 300 Chicago, 143 chicken(s), 103, 113, 142, 270, 271 China, 97, 143, 195, 203, 204, 227, 229, 299, 331 Chinese, 207 chiral center, 309 chirality, xiv, 307, 309, 318, 322, 340 chitosan, 134, 205 chloride, 26, 50, 87, 326 chlorination, 266, 285 chlorine, 130, 132, 266 chlorophenols, 131 cholesterol, 23, 173, 309 cholinesterase, 23 chromatographic technique, 268, 270, 273 chromatography, 72, 87, 111, 173, 176, 222, 224, 268, 270, 272, 273, 278, 279, 280 chromatography analysis, 176 chromosome, 212, 239 chronic lymphocytic leukemia, 86 chronic obstructive pulmonary disease, 332 chronic pain, 313 chymotrypsin, 33
Index circulation, 173 classes, 61, 103, 130, 156, 190, 264 classification, 103, 220 clean technology, 115, 181 cleaning, 121, 269, 271 cleavage, 53, 55, 62, 78, 102, 114, 133, 163, 327, 331 clone, 213 cloning, 51, 86, 212, 213, 214, 217, 224, 225, 260, 265, 266, 330 closure, 58, 59 clusters, 96 coagulation, 130 coatings, 169, 246 cocoa butter, 169, 170, 171, 178, 184 coconut, 135, 166 coding, 212, 213, 233 codon(s), xii, 230, 231, 239, 240 coenzyme, 196, 260 coffee, 105, 115, 116, 136, 139, 148, 149, 152 collagen, 205 colloid particles, 31 colloidal particles, 24, 30, 31 colloids, 34 combined effect, 289 commodity, 115 communication, 262, 263, 285 communication systems, 263 community, 213, 262, 284 compatibility, 171 competition, 290, 291 complement, 113 complications, 294 components, xiii, 31, 36, 87, 103, 173, 235, 242, 245, 251, 267, 287, 288, 336 composites, 29, 34 composition, 4, 6, 16, 18, 28, 33, 101, 102, 115, 116, 117, 118, 122, 130, 139, 142, 167, 168, 171, 270, 283 composting, 226 compounds, viii, ix, xiv, 8, 30, 47, 48, 49, 65, 74, 84, 95, 96, 97, 101, 107, 109, 111, 112, 115, 116, 121, 130, 131, 132, 133, 136, 138, 140, 141, 143, 145, 155, 163, 170, 171, 173, 174, 185, 188, 197, 215, 220, 222, 231, 244, 249, 257, 260, 262, 263, 266, 268, 269, 273, 275, 285, 290, 291, 307, 308, 309, 315, 318, 320, 322, 329, 331, 333, 336, 338, 339, 342 compressibility, 270 computers, 189
349
computing, 288 concentrates, 157, 176 concentration, xiii, 11, 22, 26, 80, 98, 100, 101, 104, 117, 119, 122, 124, 125, 126, 127, 128, 132, 133, 134, 135, 140, 149, 166, 172, 173, 180, 190, 197, 200, 201, 217, 234, 235, 236, 237, 239, 241, 246, 247, 248, 250, 253, 261, 266, 267, 271, 272, 273, 281, 282, 283, 285, 287, 289, 290, 291, 292, 293, 296, 297, 298, 299, 326 condensation, 49, 196, 333 conducting polymers, xiii, 22, 36, 245, 246, 247, 251, 255 conductivity, xiii, 245, 246, 252, 253, 255 configuration, 3, 49, 63, 246, 312, 313, 314, 327 congestive heart failure, 310 conservation, ix, 155 construction, 6, 9, 10, 17, 21, 28, 166, 192 consumption, 116, 117, 118, 119, 127, 130, 165, 286, 289, 290 contaminant(s), 49, 50, 51, 64, 89, 121, 130, 268, 271, 289, 290 contamination, 3, 52, 68, 98, 118, 132 control, vii, 1, 3, 4, 8, 9, 36, 48, 49, 100, 111, 113, 114, 121, 123, 126, 127, 130, 134, 143, 150, 156, 160, 164, 179, 182, 199, 214, 234, 237, 246, 248, 250, 252, 262, 269, 275, 309 conversion, vii, 65, 70, 72, 74, 83, 100, 121, 143, 156, 166, 172, 173, 206, 214, 235, 241, 253, 282, 285, 296, 313, 316, 327, 329, 330, 342 conversion rate, 172 cooking, 115 cooling, 269 copolymers, 257 copper, 109, 131 coral reefs, 260 corn, 98, 103, 104, 138, 142 cortex, 105 cosmetics, 166, 271 costs, vii, viii, 1, 47, 48, 51, 64, 95, 107, 111, 136, 165, 201, 267, 269, 270 cotton, 130 coupling, 142, 266 covalent bond, 3, 25, 256, 283, 284 covalent bonding, 3, 25, 284 coverage, 35, 290 covering, 31, 180, 260 critical micellar concentration (CMC), 246, 247, 248, 253 cross-linked, x, 64, 69, 195, 196, 197, 200, 201, 202, 203, 204, 205, 206
Index
350
crude oil, 133 crystal structure(s), 55, 57, 58, 60, 189 crystalline, 32, 231 crystallization, 72 crystallographic studies, 55 crystals, 27, 31, 107 cultivation, 100, 101, 102, 105, 113, 115, 117, 122, 143, 146, 221, 242, 260, 261, 263, 264, 266, 269, 271, 275, 279, 337 cultivation conditions, 100, 143 culture, ix, xiii, 95, 97, 98, 99, 101, 105, 108, 122, 123, 147, 176, 206, 212, 221, 224, 228, 263, 264, 265, 271, 274, 279, 281, 284, 285, 291, 292, 294, 296, 297, 298, 299 culture conditions, 99, 105, 123, 176, 228 cyanide, 117, 118, 196, 292, 332 cyanobacteria, 309 cycles, x, 3, 64, 72, 187, 188, 200, 203 cycling, 274 Cylindrocarpon, x, 195, 198 cystic fibrosis, 169, 182 cytidine, viii, 48, 54, 70, 77, 84 cytochrome, 17, 23, 25, 35, 320 cytometry, 35 cytoplasm, 111, 239 cytosine, 73, 74 Czech Republic, 155
D dairies, 96, 119 dairy, 119, 121, 123, 124, 125, 126, 127, 128, 138, 140, 141, 144, 145, 146, 147, 148, 151, 152, 169, 271 dairy products, 169 d-amino acids, 330, 342 data set, 190 database, 179 decay, 151 decomposition, 33, 34, 35, 116, 181 decontamination, 132 deep-sea hydrothermal vents, 275 defects, 12 defense, 105 deficiency, 337 definition, 130, 284 deformation, 14 degenerate, 189 degradation, ix, 17, 35, 48, 96, 105, 107, 109, 112, 113, 114, 115, 116, 118, 120, 121, 122, 123, 127,
128, 130, 131, 132, 133, 134, 135, 139, 141, 143, 144, 146, 148, 150, 151, 153, 190, 197, 217, 221, 226, 236, 244, 263, 264, 271, 275, 286, 290, 291, 328, 331, 342 degradation process, 128 degradation rate, 236, 290 dehydrogenases, 309 delivery, 171 demand, xiv, 19, 21, 102, 119, 122, 123, 126, 130, 222, 260, 265, 307 denaturation, xi, 5, 15, 114, 209, 211, 216, 269 denitrification, 300 density, 8, 10, 14, 27, 164, 214, 241, 244, 262, 263, 275 deoxyribose, 53, 55, 57, 60, 62, 72, 84, 333 deposition, 4, 5, 6, 8, 9, 11, 12, 13, 15, 16, 17, 18, 22, 23, 24, 25, 26, 27, 28, 29, 31, 32, 34, 36 depreciation, 165 depression, 322 derivatives, viii, ix, 48, 63, 72, 77, 133, 155, 159, 180, 196, 256, 257, 291, 313, 314, 324, 329, 342 desiccation, 210 desorption, 69, 290 detachment, xiv, 16, 281, 283, 285, 286, 287, 289, 291, 297, 298, 299 detection, 22, 30, 34, 132, 151, 177, 182, 223 detergents, 159, 231, 241, 264, 265 detoxification, viii, 95, 96, 105, 110, 111, 112, 115, 116, 117, 118, 136, 137, 139, 145, 271, 320 developed countries, 160 dialysis, 284 diastereoisomer, xv, 308 dichloroethane, 291 dielectric constant, 164 diesel fuel, 170 diet, 160, 171, 337 diffusion, xiii, 28, 33, 35, 51, 64, 99, 109, 140, 174, 200, 205, 231, 262, 270, 281, 289, 292, 293, 294 diffusion process, 270 diffusivities, 164, 165, 293, 294, 296 diffusivity, 99, 282, 296 digestibility, 104, 113, 115, 117, 141 digestion, 100, 103, 114, 118, 123, 124, 137, 139, 140, 143, 146, 149, 152, 169, 264, 270 digestive tract, 160 dimer, 57 dimethylformamide, 191 direct measure, 8 directives, 48 discrimination, 163
Index discs, 286 disinfection, 285 disorder, 175, 311 dispersion, 25 displacement, 269 dissociation, 11, 52, 66 dissolved oxygen, 127 distillation, 166, 325 distilled water, 11, 115, 247, 248 distribution, xi, 13, 19, 167, 176, 209, 269, 270, 286, 289, 293, 294, 298 diversity, ix, x, xi, xiii, 102, 123, 187, 189, 190, 207, 209, 210, 213, 214, 219, 220, 221, 222, 225, 226, 228, 259, 267, 274, 277 division, 13 D-lactonohydrolase, x, 195, 199, 200 DMF, 68, 76, 77, 79 DNA, viii, ix, 21, 24, 47, 61, 84, 86, 87, 187, 188, 189, 190, 191, 192, 199, 212, 213, 214, 260, 262, 265, 266, 269, 271, 276, 277 DNA polymerase, 24, 188, 261, 265, 267, 276 DNase, 87 docosahexaenoic acid (DHA), 156, 158, 163, 167, 168, 169, 176, 179, 180, 183 dodecyl diphenyloxide disulphonate (DODD), xiii, 245, 247, 248, 253, 254, 255 domain structure, 12, 19 donors, 15, 63, 74, 85, 86, 87, 168 dopamine, 156 doping, 246 dosage, 52 double bonds, 120, 156, 162, 163, 169, 171, 309 d-pantoic acid (D-PA), x, 195, 196, 197, 198, 199, 200, 202, 203 d-pantolactone (D-PL), x, 195, 196, 197, 198, 199, 200, 205 Drosophila, 86 drug use, 86, 331 drugs, xiv, 48, 89, 222, 226, 267, 307, 313, 314, 323, 328, 329, 331, 338, 339 drying, 13, 117 DSM, 123, 342 dyeing, 131, 150 dyes, 107, 130, 131, 132, 133, 135, 137, 146, 147, 148, 149, 151, 153 dyspepsia, 316
E earth, 220, 260
351
eating disorders, 313 ecology, 214, 226, 277 ecosystem, 96, 261, 276 effluent(s), 97, 98, 107, 109, 119, 121, 122, 123, 124, 125, 127, 128, 129, 130, 132, 133, 134, 136, 138, 139, 140, 144, 146, 147, 148, 150, 151, 152, 153, 273, 286, 301, 302 egg, 118, 270 Egypt, 226, 227 eicosapentaenoic acid, 156, 163, 175, 176, 179, 182 electric charge, 7 electric field, 270 electrical conductivity, xiii, 245, 246, 248, 252, 255 electrocatalytic, 36 electrochemical reaction, 22, 30 electrochemistry, 257 electrodes, 22, 27 electrolytes, 246 electromagnetic, 246 electron(s), 5, 15, 21, 22, 28, 36, 55, 109 electron density, 21 electrophoresis, 111 electrostatic interactions, 7, 16, 17, 283 emission, 96 employment, ix, x, 26, 96, 97, 113, 132, 136, 196 emulsification, 166 enantiomer(s), xiv, xv, 197, 307, 308, 309, 319, 323, 325, 330, 336, 337, 342 enantioselective synthesis, xiv, 307, 339, 340 encapsulation, 3, 31, 32, 33, 34, 252 encoding, 86, 212, 216, 233, 244, 267, 341 endonuclease, 24 endotoxins, 271 energy, ix, 15, 28, 53, 55, 87, 100, 113, 119, 148, 155, 159, 165, 169, 171, 203, 204, 242, 271 energy consumption, 100 energy supply, 113 energy transfer, 15 entrapment, vii, xiii, 1, 3, 200, 281, 283, 284, 295, 296 entropy, 7 environment, 96, 97, 101, 118, 173, 197, 221, 231, 246, 263 environmental conditions, xiii, 102, 117, 131, 143, 259, 261, 262 environmental effects, 121 environmental impact, viii, 47, 48 Environmental Protection Agency (EPA), 156, 158, 167, 168, 169, 191 environmental technology, 98, 102, 136
Index
352
enzymatic activity, 5, 15, 25, 33, 68, 107, 200, 249 enzymatic transformation, 329 enzyme immobilization, 3, 66, 89 enzyme induction, 337 enzyme interaction, 21 epoxy, 68, 69 epoxy resins, 69 equilibrium, 11, 21, 63, 70, 80, 81, 87, 172, 249, 290 equipment, 17, 18, 121, 165, 205, 238, 270, 301 Escherichia coli (E. coli), xii, 54, 55, 57, 58, 60, 64, 65, 66, 67, 69, 71, 87, 88, 191, 211, 212, 229, 230, 231, 233, 236, 237, 240, 242, 243, 244, 263, 283, 314, 315, 316, 337, 342, 343, 344 essential fatty acids, 156 ester bonds, ix, 105, 155, 163 esterification, ix, 100, 155, 158, 160, 161, 167, 169, 170, 172, 176, 177, 180, 181, 182, 184, 185 ester(s), ix, 63, 75, 76, 79, 83, 87, 88, 100, 103, 105, 121, 146, 155, 156, 161, 163, 167, 168, 170, 172, 173, 174, 180, 181, 184, 185, 197, 205, 309, 313, 324, 327, 339, 341 ethanol, 33, 79, 170, 172, 184, 185, 292, 323 ethers, 320 ethylene, 316, 340 EU, 48 eukaryotes, 100, 221 Europe, 84 European Community, 267 evening, 157, 168, 176, 182 evidence, 57, 58, 127, 148, 156 evolution, ix, 124, 187, 188, 189, 190, 191, 192, 193, 199, 211, 214, 329 excision, 239 excitation, 19 exclusion, 231 excretion, 104 exonuclease, 276 experimental condition, 52, 167, 170 experimental design, 145 exploitation, 260, 262 exposure, 130, 327 extraction, x, 64, 107, 108, 111, 173, 174, 185, 195, 199 extremophiles, 210, 220, 226, 261, 264
F F. solani, 198 fabrication, 3, 4, 24, 25, 26, 28, 30, 31, 34, 35, 36 failure, 120
family, 54, 55, 57, 60, 100, 160, 162, 226, 275, 322 family members, 55 farmers, 147 fat, 97, 100, 116, 119, 121, 122, 124, 125, 127, 128, 138, 141, 144, 146, 148, 165, 166, 170, 172, 178, 181 fatty acids, ix, 7, 11, 17, 100, 101, 120, 121, 128, 137, 143, 149, 150, 155, 156, 157, 158, 159, 160, 163, 165, 166, 168, 169, 170, 171, 173, 175, 176, 182, 183, 185, 263 feces, 147 feedback, 9, 12, 16, 18, 19, 20 fermentation, ix, x, xiii, 84, 90, 95, 96, 97, 98, 99, 101, 102, 103, 104, 105, 108, 111, 115, 116, 117, 118, 123, 124, 136, 137, 138, 139, 140, 141, 142, 143, 144, 145, 146, 147, 148, 149, 150, 151, 152, 196, 261, 263, 267, 269, 271, 281, 283, 294, 295, 297, 298, 331 fermentation broth, xiii, 261, 269, 271, 281 ferrous ion, 146 fiber membranes, 257 fibers, 130, 256 fibrin, 279 fibrosis, 169, 313 filament, 115, 150 film formation, xiii, 245, 250, 255 film thickness, 23, 289 film(s), vii, xiii, 1, 4, 5, 6, 11, 13, 14, 15, 17, 22, 23, 24, 26, 27, 28, 29, 30, 32, 33, 35, 36, 245, 250, 252, 255, 289 filters, 119 filtration, 52, 111, 142, 148, 199, 201, 203, 224 fish oil, 156, 158, 163, 172, 175, 179, 180 fission, 332 fixation, xiii, 281, 284 flammability, 164 flavonoids, 336 flavor, 84, 159 flexibility, 81, 160, 171, 189, 269 floating, 11, 120, 285 flocculation, 130, 283, 284 flora, 122, 220 flotation, 119, 121, 124 fluctuations, x, 195, 199, 231 fluid, 165, 172, 173, 174, 177, 184, 185, 202, 285, 289, 299 fluid extract, 173, 174, 177, 184, 185 fluidized bed, 285 fluorescence, 19, 20, 33 fluoxetine, 313, 339
Index focusing, 263, 264, 267, 270, 271 folic acid, 326 food, 2, 49, 77, 82, 84, 89, 97, 101, 105, 108, 115, 117, 119, 120, 122, 124, 150, 151, 159, 166, 168, 169, 170, 172, 173, 180, 188, 197, 213, 221, 223, 276, 285, 286, 322, 333, 335 food additives, 49, 84, 166, 180, 223 food industry, 82, 101, 120, 124, 151, 159, 173, 197, 333 food products, 119, 172, 335 formaldehyde, 33, 34, 196, 332 formamide, 215 Fourier transform infrared spectroscopy, 28 fragmentation, 138, 192 France, 148 freezing, 107, 110 Friedmann, 43 fuel, viii, 2, 144 fulfillment, 63 functional analysis, 213 functionalization, 82, 159 funding, 175 fungus(i), x, 96, 97, 98, 99, 100, 102, 103, 104, 105, 108, 111, 115, 116, 117, 124, 131, 132, 135, 138, 139, 140, 142, 144, 146, 149, 151, 152, 153, 158, 160, 176, 178, 195, 198, 199, 203, 260, 261, 262, 315 Fusarium, x, 195, 198, 199, 205, 206, 207, 326, 342 Fusarium oxysporum, 198, 199, 205, 206, 207, 326
G gases, 99, 165, 184, 210 gastroesophageal reflux disease, 316 gel, xiii, 34, 111, 200, 205, 224, 242, 281, 283, 284, 292, 293, 294, 295, 296, 298 gelation, 284 gene(s), xii, 86, 104, 188, 189, 190, 192, 199, 205, 210, 212, 213, 214, 216, 217, 222, 224, 225, 228, 229, 231, 233, 239, 244, 260, 262, 263, 266, 267, 269, 277, 289, 341 gene amplification, 212 gene expression, 213, 262 generalization, 242 generation, ix, xii, 96, 100, 111, 112, 130, 136, 165, 168, 192, 211, 220, 260, 265, 289, 338 genetic blueprint, 213 genetic diversity, 284 genetically modified organisms, 97 genome, 213, 218
353
genomics, 215, 217, 225, 267, 277 Georgia, 150, 229 Germany, 272, 286, 301 Gibberella, x, 195, 198 glass(es), 26, 184, 269, 283 glucoamylase, 35, 151, 264 glucose, xii, 22, 25, 30, 31, 34, 35, 36, 101, 102, 105, 131, 153, 205, 230, 236, 237, 238, 239, 241, 244, 273, 309, 310, 319 glucose oxidase, 25, 30, 31, 34, 35, 36 glucosidases, 105 glutamate, 23 glutaraldehyde, x, 64, 195, 196, 200, 201, 202, 203, 204, 205, 206, 284 glutathione, 24 glycerol, ix, 100, 120, 155, 158, 159, 160, 163, 165, 166, 167, 168, 170, 171, 178, 181, 182, 215, 309, 324 glycoproteins, 109 glycoside, 49 glycosylation, viii, 47, 49, 50, 63, 64, 72, 75, 83 glycosyl-transferring enzymes, viii, 48, 67 glyphosate, 190 goals, 287 government, 130 grains, 103 gram-negative, xii, 230, 231, 232, 271, 275, 280, 284 gram-negative bacteria, 242, 263 gram-positive bacteria, 263 groups, viii, x, xiv, 3, 4, 6, 10, 11, 12, 13, 15, 16, 17, 19, 20, 47, 49, 58, 65, 68, 69, 75, 78, 84, 89, 102, 107, 133, 163, 169, 176, 195, 201, 224, 247, 253, 270, 281, 283, 285, 327 growth, ix, xiii, xiv, 27, 33, 34, 95, 98, 100, 103, 111, 113, 115, 116, 118, 123, 141, 175, 188, 214, 221, 225, 241, 243, 261, 263, 281, 282, 283, 285, 286, 287, 289, 291, 293, 294, 295, 297, 299, 322, 334 growth rate, 214, 282, 285, 289, 295, 299 guanine, 54, 67, 70, 71, 72, 74 guanosine, viii, 48, 64, 69, 72, 78, 84 guidance, 90 guidelines, 48, 172 Gujarat, xi, xii, 209, 211, 214, 215, 216, 218, 220
H habitat, 99, 220, 260 half-life, 68, 201, 265, 326 haloalkaliphiles, xi, 215, 219
354
Index
haloalkaliphilic, xi, 209, 211, 214, 215, 223, 228 halogen, 266 halogenation, 266 halophiles, xi, 210, 219 halophilic, 213, 214, 216, 217, 218, 222 harmonization, 48 health, 167, 168, 175 heart disease, 337 heart rate, 310 heat, 68, 98, 110, 269, 273, 276, 280, 309, 324 heating, 121, 198 heavy metals, 130 heme, 109, 266 hemoglobin, 29, 30 Heparin, 279 heptane, 321, 335 herpes, 86, 279 herpes simplex, 86 heterogeneity, xi, 100, 210, 211, 221, 284 heterogeneous catalysis, 292 heterotrophic, 271 hexafluorophosphate, xiii, 174, 245, 252 hexane, 215, 335 high fat, 121, 122, 123, 124, 127, 128, 139, 145 histochemistry, 109 HIV, 49, 74, 75, 279, 314, 319, 339 HIV-1, 74 hog, 172 holotoxin, 111 homogeneity, 9, 12 homologous genes, 189 homopolymers, 257 hormone, 226 horseradish peroxidase(s) (HRP), xiii, 109, 132, 133, 134, 149, 245, 246, 247, 248, 249, 251, 252 host, 212, 260, 263, 285 HPLC, 81, 167, 182 human immunodeficiency virus, 279, 339 human kinase, 85, 86 human milk, 84, 171 hybrid, 29, 119, 185 hydrocarbons, 131, 216 hydrochloric acid, 247 hydrofluoric acid, 29 hydrogen, 22, 29, 54, 58, 59, 60, 61, 62, 63, 107, 109, 117, 131, 134, 135, 146, 150, 156, 247, 248, 286, 309, 331, 333 hydrogen atoms, 156 hydrogen bonds, 61, 62 hydrogen cyanide, 117, 331
hydrogen peroxide, 22, 29, 107, 109, 131, 134, 135, 146, 150, 247, 248 hydrolases, viii, x, 48, 53, 75, 79, 80, 89, 96, 103, 160, 184, 195, 275, 321, 322, 329, 342 hydrolysates, 84, 141 hydrolysis, ix, 21, 50, 75, 76, 77, 78, 79, 81, 82, 83, 84, 87, 89, 99, 100, 102, 105, 114, 120, 122, 123, 124, 125, 126, 127, 128, 129, 136, 139, 143, 145, 146, 150, 152, 155, 158, 159, 160, 161, 163, 165, 166, 167, 171, 172, 176, 177, 180, 181, 182, 184, 197, 198, 199, 200, 201, 202, 207, 215, 241, 279, 322, 323, 324, 326, 327, 329, 330, 341, 342 hydrolysis kinetics, 200 hydroperoxides, 315, 340 hydrophilic, xii, 7, 17, 21, 62, 166, 230, 231, 233, 236 hydrophilic groups, 21 hydrophobic, xii, 5, 7, 11, 15, 17, 34, 52, 62, 68, 80, 81, 89, 121, 145, 230, 231, 233, 234, 235, 236, 243, 253 hydrophobic groups, 15 hydrophobic interactions, 11, 34 hydrophobicity, xii, 184, 229 hydroxyapatite, 205 hydroxyl, 50, 60, 63, 65, 76, 77, 81, 82, 83, 84, 158, 171, 319 hydroxyl groups, 83, 84, 158, 171 hyperglycemia, 311 hypothesis, 99, 131
I ibuprofen, 336 identification, 2, 151, 153, 189, 228, 260, 263, 264, 288 identity, 54, 189, 212, 266 images, xiii, 245, 302 immersion, 9 immobilization, vii, viii, x, xiii, xiv, 1, 3, 4, 17, 29, 30, 36, 48, 52, 53, 66, 67, 68, 69, 75, 81, 85, 89, 195, 199, 200, 246, 251, 281, 282, 283, 284, 294, 295, 299, 305 immobilized enzymes, vii, 1, 3, 28, 51, 52, 68, 77, 80, 85 immunoglobulin, 30 immunological, 109 immunosuppressive agent, viii, 47 impurities, 29, 48, 90, 269 in situ, 71, 128, 201, 289 in situ hybridization, 289
Index in vitro, 188, 191, 192, 242 in vivo, 55, 243, 335 incidence, 20 inclusion, 31, 121 incubation period, 122 India, xi, xii, 209, 211, 214, 215, 216, 218, 219, 220, 225, 226, 259 Indonesia, 97 induction, xiv, 105, 275, 308 industrial application, 2, 4, 22, 31, 89, 97, 107, 160, 168, 169, 190, 197, 201, 204, 211, 213, 222, 223, 273, 282, 284, 299 industrial production, 96, 100, 160, 175, 329 industrial sectors, 2 industrial wastes, 116, 136 industry, vii, ix, xiii, 1, 2, 96, 97, 102, 107, 112, 119, 124, 128, 129, 131, 135, 138, 140, 143, 144, 146, 152, 172, 173, 174, 187, 218, 225, 259, 264, 267, 286, 309, 318, 327, 333 infancy, xi, 219 infants, 84 infection, 74, 263, 279 inflammatory disease, 171 inhibition, 29, 119, 128, 132, 143, 144, 188, 190, 234, 295, 297 inhibitor, 273, 283, 290, 295, 296, 309, 315, 316, 319, 331, 339 inhibitory effect, 120 inoculation, 116, 285 inoculum, 98, 116, 118, 122, 123, 145, 148, 151 inositol, 103 insertion, xii, 230, 231, 233, 239, 240 insight, 214, 221, 260, 263, 268, 289 instability, 51, 67, 166 instruments, 8, 27 integrity, 29 intensity, 28, 35 interaction process, 21, 22 interaction(s), 3, 5, 6, 11, 14, 15, 16, 17, 18, 19, 20, 21, 25, 35, 54, 60, 63, 68, 80, 81, 138, 160, 184, 193, 213, 235, 254, 261, 262, 263, 272, 279, 283, 284, 289, 328 interface, viii, 2, 4, 5, 6, 7, 8, 9, 10, 11, 12, 15, 17, 18, 19, 20, 21, 27, 28, 160, 162, 251, 253, 257, 271, 285, 294, 298 interference, 113, 137 interpretation, 10, 19 interval, 134 invertebrates, 261 investment, 102
355
ion channels, 175 ion exchangers, 270 ionic liquid(s), xiii, 159, 164, 165, 174, 177, 181, 245 ions, 11, 16, 21, 31, 165, 174, 235, 253, 285 iron, 109, 162, 261, 266, 274 iron transport, 274 irradiation, 151 isolation, x, 84, 111, 149, 195, 199, 213, 221, 224, 230, 260, 265, 270, 273, 274, 278, 279, 283, 327 isomers, 50, 72, 89, 120, 163, 196 Italy, 1, 47, 142
J Japan, 84, 97, 122, 165, 166, 203, 207 Jordan, 123, 143
K K+, 210 Kenya, 171 keratin, 103, 112, 113, 114, 141, 147, 150 kernel, 150, 179, 181 ketones, 322, 331, 339, 341 kidney, 324 kinase(s), 84, 85, 86, 87, 212 kinetic constants, 299 kinetic curves, 298 kinetic model, 139, 294 kinetic studies, 161 kinetics, xiv, 22, 123, 160, 182, 270, 281, 297 Kuwait, 225
L labeling, 172, 184, 265 labor, 201 lack of control, vii, 1, 3 lactic acid, 61, 118, 142, 144, 294, 295, 298 lactic acid bacteria, 61, 142, 144 Lactobacillus, 57, 61, 62, 105, 118, 147, 295, 298 lactones, 121, 199, 207, 263, 341 lakes, x, 209, 210 land, 119 Langmuir-Blodgett, viii, 2, 4, 7, 11, 23, 39, 43, 44 language, 275 large intestine, 171 laser, 20, 35
356
Index
latex, 31 Latin America, 117 laundry, 223 L-carnitine, xii, 230, 240, 241, 244, 319, 337 leachate, 286 lead, ix, x, 107, 130, 131, 156, 159, 164, 187, 195, 201, 239, 242, 260, 269, 299 leakage, x, xiii, 196, 201, 281, 282, 297, 298, 299 lecithin, 21 lectin, 111, 273, 280 legislation, 130 legumes, 103 Lentinula edodes, 147 leucine, 329 Lewis acids, 249 liberation, 269 life sciences, 188 ligand(s), 273, 274 light scattering, 34 lignin, 29, 30, 107, 131, 132, 133, 138, 140, 142, 143, 149, 151, 190, 192 likelihood, 84 limitation, vii, xv, 1, 3, 21, 28, 121, 233, 241, 242, 308 linkage, 71, 89, 163, 200, 239, 260 links, 201 lipid metabolism, 100 lipid(s), 6, 16, 18, 19, 20, 21, 23, 100, 101, 120, 123, 128, 149, 150, 152, 159, 160, 168, 169, 177, 178, 179, 182, 183, 337 lipolysis, 163, 182 lipopolysaccharide(s) (LPS), xii, 148, 150, 229, 231, 233, 234, 235, 237, 238 lipoprotein(s), 88, 100, 241, 244 liquefaction, 120, 223, 264 liquid chromatography, 182, 214 liquid interfaces, 27 liquid phase, 98, 99, 251, 252, 286, 288, 294 liquids, 160, 165, 181 literature, xi, 6, 64, 80, 85, 89, 99, 101, 107, 115, 119, 172, 209, 219, 221, 223, 224, 291, 299, 318 lithium, 87 liver, 158, 167, 172, 184, 271, 313 livestock, 118 localization, 21 location, xv, 145, 163, 169, 239, 308 London, 39, 146, 175, 226, 256, 278, 279, 304 loss of appetite, 313 Louisiana, 45 low temperatures, 264, 265
lubricants, 170 luciferase, 23, 30, 263 lysine, 60, 113, 141, 316 lysis, 107, 269 lysozyme, 267, 270, 278
M machine learning, 193 machinery, xiii, 259 macromolecules, 24, 34, 69, 210, 221 macroorganisms, xiii, 259, 260, 261 magnesium, 283 magnet, 10 magnetic field, 31 magnetic properties, 31, 34, 36 magnetite, 34 malate dehydrogenase, 24 Malaysia, 171 males, 175 maltose, 223, 225 mammal, 85 mammalian cells, 111, 156 mammalian tissues, 271 management, 149 manganese, 30, 108, 109, 131, 135, 143, 151 manipulation, 4, 100, 111, 164, 190, 324 manufacturing, xiii, 51, 231, 259, 260, 270 marine environment, 221, 222 market, xiv, 48, 80, 100, 102, 104, 107, 191, 203, 265, 307, 313, 333 market value, 333 marketing, 48 marsh, 225 mass spectrometry, 224, 227 mass transfer process, 99 Massachusetts, 243 mast cell, 111 matrix, 52, 99, 171, 200, 262, 270, 273, 283, 284, 285 maturation, 107, 285, 287 maximum specific growth rate, 295 meals, 332 measurement, 8, 21, 32, 147, 237, 248, 253 measures, 8, 35 meat, 96, 119, 150 mechanical properties, 35, 299 media, 4, 29, 80, 102, 113, 134, 137, 145, 147, 159, 161, 164, 165, 174, 177, 181, 215, 246, 249, 251, 252, 253, 261, 269, 285, 286, 292, 322
Index medication, 313 medicine, ix, 3, 155, 320 melamine, 33, 34 melting, 171, 178 membrane permeability, xii, 229, 230, 233, 235, 239, 241, 243, 244 membranes, 6, 21, 29, 52, 174, 231, 237, 268, 270, 284, 292, 295, 298 mental illness, 156, 175 metabolic pathways, 214, 337 metabolism, xii, 2, 100, 131, 133, 168, 205, 230, 231, 244, 271, 279, 287, 299 metabolites, xi, 131, 156, 219, 221, 222, 225, 227, 260, 262, 263, 266, 274, 277 metabolizing, 242 metagenomics, xi, 210, 213, 214 metal ions, 11, 12, 235 metalloproteinase, 103, 265, 276 metals, 210 methane, 120, 121, 123, 128, 140 methanol, 215 methionine, 60, 113, 141 methyl group, 163 methylation, 336 methylene, 133, 135, 141, 142, 144, 150, 151, 162 Mg2+, 103 micelles, 19, 20, 246, 253, 256, 311 microbial, 87, 93, 137, 138, 139, 143, 146, 150, 177, 179, 205, 216, 217, 221, 231, 243, 254, 255, 258, 275, 277, 281, 284, 300, 309, 313, 318, 319, 331, 339, 343 microbial cells, xv, 205, 206, 282, 283, 284, 290, 291, 293, 299, 308, 309, 337 microbial community(ies), 213, 267, 269, 275 microelectronics, 246 microenvironment, 28 micrometer, 164 microorganism(s), xiv, 51, 61, 64, 68, 76, 97, 98, 99, 100, 101, 102, 103, 104, 105, 108, 113, 114, 115, 117, 118, 120, 121, 123, 127, 132, 146, 147, 162, 198, 199, 206, 207, 210, 211, 213, 214, 215, 220, 221, 223, 224, 228, 260, 261, 262, 263, 264, 271, 273, 274, 275, 276, 278, 284, 285, 288, 307, 309, 311, 314, 316, 320, 322, 325, 331, 336, 340 microscopy, xiii, 12, 19, 20, 28, 33, 35, 245, 250, 289 microspheres, 33, 36 microstructure, 205 migraine, 320 migration, 170, 178
357
milk, 84, 119, 148 minerals, 117 mining, 286 Ministry of Education, 175, 195, 229 mixing, 31, 164, 167, 286 mobility, vii, 1, 34 model system, xi, 29, 210, 211 modeling, xiv, 189, 192, 281, 283, 287, 288, 289, 291, 294, 295, 298 models, 59, 190, 220, 287, 288, 289, 294, 295, 299 moderate activity, 79 modules, 271 modulus, 27, 282, 293 moieties, 167 moisture, 100, 102, 103, 116, 141, 173, 185 molasses, 104, 113, 116 molecular beam, 23 molecular beam epitaxy, 23 molecular biology, xiii, 259, 265, 267, 274, 289, 341 molecular mass, 109, 146 molecular oxygen, 106, 107, 131 molecular structure, 6 molecular weight, 63, 213, 253 molecules, vii, ix, xv, 1, 2, 3, 4, 5, 6, 7, 8, 10, 11, 13, 15, 16, 17, 19, 20, 21, 22, 23, 25, 26, 27, 28, 29, 30, 31, 32, 33, 36, 81, 109, 119, 127, 130, 131, 155, 160, 165, 167, 168, 172, 188, 224, 231, 235, 236, 246, 253, 262, 266, 285, 290, 293, 308 momentum, 214 monoamine oxidase, 23 monoclonal antibodies, 271 monograph, 90 monolayer, 4, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22 monomer(s), 109, 148, 212, 213, 246, 247, 249, 251 monomer molecules, 246 mononucleotides, 49 Montana, 340 mood, 156 morphology, 19, 28, 148, 210, 221, 248, 289 motion, 11, 12, 13, 14, 16, 17, 18 movement, 58, 59 mRNA, 214 mucosa, 72, 273 multilayer films, 28, 29, 32, 35 multilayered structure, 25, 26, 35, 36 multiplicity, 100 mushrooms, 107, 108, 135 mutagenesis, ix, 187, 188, 189, 190, 191, 192, 225, 239, 329
Index
358
mutant, xii, 176, 191, 199, 229, 230, 231, 233, 234, 235, 237, 239, 241, 243 mutant cells, xii, 230, 241 mutation(s), ix, x, xii, 187, 188, 189, 190, 191, 192, 195, 199, 229, 233, 231, 233, 234, 235, 237, 238, 239, 241, 242, 243, 244, 276 mycelium, 98, 326 Mycobacterium, 226 myoglobin, 25
N NaCl, 29, 33, 213, 215, 271, 272 NAD, 237, 238, 310 nanocomposites, 205, 256, 257 nanofabrication, 3, 4 nanometers, 25 nanoparticles, 24, 29, 31, 34, 36 nanoreactors, 30, 31 nanostructured materials, 3 nanostructures, 4 nanotechnology, vii, 1, 2, 3 native protein conformation, 201 natural environment, 112, 213, 242, 266, 267, 309 natural habitats, 262 natural polymers, 170 neural systems, 156 neuronal excitability, 320 neurons, 156 neurotransmitter, 313 New York, 39, 41, 43, 90, 138, 141, 176, 179, 242, 243, 257, 274, 277, 278, 279, 300, 304, 338 New Zealand, 139 next generation, 3, 37 Nigeria, 147 nitrate, 289 nitrification, 285 nitrilase, 327 nitrile hydratase, 327 nitrobenzene, 291 nitrogen, 13, 27, 53, 59, 65, 97, 101, 103, 115, 116, 334 Nobel Prize, 109 non-conventional, xii, 159, 164, 220 novelty, 222 nuclear magnetic resonance (NMR), 167, 182 nucleic acid, 84, 93, 191 nucleosides, viii, 47, 48, 49, 50, 51, 53, 54, 61, 63, 64, 65, 70, 71, 72, 74, 75, 76, 77, 78, 79, 80, 81, 82, 83, 84, 85, 86, 87, 89, 90
nucleotide sequence, 212, 240, 244 nucleotides, viii, 47, 48, 49, 50, 51, 53, 84, 85, 87, 212, 326 nucleus(i), 5, 34 nutraceutical, 168 nutrients, 98, 99, 100, 103, 115, 119, 231, 262, 285, 287, 294 nutrition, 118, 150, 152, 160 nuts, 122
O obesity, 313, 329 observations, 292, 294 observed behavior, 242 obstructive lung disease, 311 oceans, xiii, 259, 260, 274 octane, 335 oil(s), ix, 80, 101, 102, 104, 110, 119, 121, 122, 125, 129, 133, 139, 142, 143, 144, 146, 147, 149, 150, 151, 152, 153, 155, 157, 158, 159, 161, 163, 164, 165, 166, 167, 168, 169, 170, 171, 172, 173, 176, 177, 178, 179, 180, 181, 182, 183, 184, 185, 214, 218, 221, 223, 224, 285 oil production, 147 oilseed, 177 Oklahoma, 214 oligosaccharide, 231 olive oil, 101, 102, 116, 123, 136, 138, 141 omega-3, 175, 176, 179 omeprazole, 316 oncology, 90 one dimension, 289 operator, 212 operon, 233 optical properties, 246 optimal performance, 271 optimization, viii, 95, 96, 112, 152, 176, 181, 206, 260 organelles, 29 organic compounds, 109, 132, 136, 286, 338, 341 organic matter, 96, 119, 123, 127, 140 organic solvent(s), 51, 52, 66, 68, 76, 151, 160, 161, 164, 165, 167, 169, 174, 177, 179, 183, 188, 205, 214, 215, 216, 217, 241, 335, 343 organism, 166, 213, 261, 263, 265, 269, 271, 322, 331 organization, viii, 2, 15, 19 orientation, 3, 5, 10, 13, 15, 16, 21, 60, 63 oscillation, 287
Index osmolality, 262 osmosis, 152 osmotic, 210, 270 osmotic pressure, 270 oxidation, 29, 33, 36, 106, 108, 109, 130, 131, 132, 133, 136, 141, 142, 144, 145, 148, 197, 206, 246, 247, 249, 271, 318, 320, 321 oxidation rate, 134 oxides, 283 oxygen, 11, 58, 59, 60, 63, 100, 101, 109, 119, 120, 122, 123, 126, 127, 130, 162, 210, 287, 289, 294, 299, 318, 327, 342 ozone, 130
P Pacific, 264 palliative, 113 palm oil, 142, 162, 171, 178, 179 pancreas, 75, 100, 172 pancreatic cancer, 156, 175 pancreatic insufficiency, 182 pantothenic acid, 199, 203, 204 parameter, 7, 10, 15, 120, 172, 217, 288 parenchyma, 105, 117, 118 parents, 190 particle collisions, 286 particles, xiii, xiv, 30, 31, 33, 35, 36, 122, 124, 146, 178, 206, 267, 281, 283, 286, 287, 289, 291, 293, 294, 297, 298, 299 partition, 164, 214 passive, 231 patents, 64, 121, 265 pathogenic, 263 pathways, 157, 213, 227, 263 Pediococcus, 233 penicillin, 17, 24, 190, 192 pepsin, 5, 141 peptic ulcer, 316 peptidase, 103 peptides, 102, 263, 265 perception, 147 performance, vii, xiii, 22, 64, 80, 120, 123, 124, 130, 132, 134, 136, 137, 143, 147, 148, 151, 152, 167, 182, 216, 260, 263, 264, 265, 266, 269, 270, 273, 281, 287, 299, 329 performers, 189 permeability, xii, 35, 229, 230, 231, 233, 234, 235, 236, 237, 238, 239, 241, 242, 244 permeation, 231
359
permittivity, 34 peroxide, 29, 109, 133, 134, 190, 247 petrochemical, 286 pH, vii, x, 2, 11, 15, 16, 23, 25, 26, 28, 32, 33, 51, 52, 66, 67, 68, 69, 70, 71, 78, 81, 87, 88, 100, 103, 105, 118, 122, 130, 131, 134, 135, 166, 173, 188, 196, 199, 200, 202, 210, 211, 213, 215, 220, 221, 223, 224, 246, 247, 248, 249, 262, 264, 265, 270, 271, 272, 283, 289, 292, 294, 334, 337, 340 pharmaceuticals, vii, 177, 221, 271, 309, 339, 340 pharmacology, 159, 173 pharmacopoeia, xiv, 307 phase transitions, 8, 19 phenol, 106, 114, 134, 136, 143, 151, 285, 291, 335 phenotype(s), xii, 230, 239, 241 phenylalanine, 316, 340 phosphatases, viii, 48, 53, 88, 105 phosphate(s), 53, 55, 57, 58, 59, 60, 63, 68, 71, 72, 76, 77, 79, 81, 82, 84, 85, 86, 87, 88, 103, 105, 135, 333, 334 phospholipids, 163, 168, 179, 180 phosphorous, 104 phosphorus, 82, 103, 104 phosphorylation, viii, 48, 53, 82, 83, 84, 85, 86, 87, 88, 89 photosynthesis, 130 phylogeny, xi, 209, 210, 215, 219, 221 physical interaction, 68 physical properties, 121, 158, 165 physical treatments, 231 physicochemical, 144 physico-chemical characteristics, 167 physico-chemical properties, 25, 165, 170 physiological factors, 115 physiology, xiii, 241, 242, 281, 294, 299 phytates, 104 phytosterols, 163, 173 Pichia pastoris, 316 piezoelectric, 27 pigs, 103, 320 placebo, 175 planning, 291 plants, 96, 100, 102, 104, 105, 119, 121, 130, 160, 162, 174, 185, 260, 271, 335 plasma, 28, 337 plasma membrane, 337 plasmid, 210, 212, 241 platinum, 333 Pleurotus ostreatus, 108, 133, 150 pneumonia, 320, 324
360
Index
Poland, 307 polarization, 21 polarized light, 20 pollutants, 29, 130, 131, 133, 149, 217, 285, 286, 290, 291 pollution, ix, 89, 96, 97, 103, 115, 119, 130, 150 polyacrylamide, 284, 295 polyaniline (PANI), xiii, 22, 245, 246, 247, 248, 251, 252, 253, 255, 256, 257 polycarbonate, 29 polyelectrolytes, 4, 25, 31, 34, 35 polyesters, 121, 146 polyethylenimine, 69, 241 polymer(s), xiii, 23, 29, 31, 35, 69, 121, 179, 211, 245, 246, 247, 248, 250, 253, 255, 256, 257, 273, 283, 288, 295 polymer film(s), 29, 255 polymer matrix, 283 polymer synthesis, 256 polymerase, 24, 191, 265, 277 polymerase chain reaction (PCR), 188, 191, 192, 199, 212, 265, 267 polymerization, xiii, 106, 107, 245, 246, 247, 248, 249, 253, 254, 255, 256, 257, 284 polymerization process, 248, 250 polymerization temperature, 248 polymerization time, 248, 249 polypeptide, 111 polyphenols, 29, 115, 116, 136 polypropylene, 179, 257 polysaccharide, 231, 279 polystyrene, 30, 34, 36 polystyrene latex, 30 polystyrenesulfonate (PSS), xiii, 26, 29, 32, 33, 34, 245, 246, 248, 250, 255 polythiophenes, 247 polyunsaturated fat, 156, 157, 158, 159, 162, 163, 166, 168, 169, 172, 175, 176, 179, 180, 182, 183 polyunsaturated fatty acids, 157, 158, 159, 166, 168, 169, 172, 175, 176, 179, 180, 182, 183 polyurethane, 151, 177, 295 polyurethane foam, 295 poly(acrylic acid) (PAA), 29, 295 pools, 121, 190 poor, 64, 115, 117, 118, 120, 122, 246, 250, 261 population, xi, 123, 189, 213, 214, 219, 251, 262, 266, 277, 285 population density, 262 porosity, 23 Portugal, 139
potassium, 81, 320 poultry, 103, 112, 113, 118, 128, 129, 150, 152, 320 power, 164, 174, 286, 288 precipitation, 31, 87, 107, 131, 246, 253, 290, 324 prediction, 226 preference, 62, 86 premature infant, 171 pressure, xiii, 6, 7, 8, 9, 10, 12, 15, 16, 18, 19, 20, 22, 35, 110, 159, 164, 165, 166, 167, 172, 173, 177, 181, 182, 188, 202, 210, 220, 259, 265, 269, 270, 286, 334 prices, 3 prions, 273 probability, 190 probe, 10, 19, 248, 267, 289 process control, 3, 51, 270 prodrugs, 49, 79 producers, 101, 105, 108, 118, 138, 171 production costs, 51, 89, 265 productivity, 88, 98, 99, 101, 105, 112, 119, 200, 203, 283, 286, 292, 298 prokaryotes, 221 promote, 118, 121, 165, 201 promoter, 49, 212 prophylaxis, 320 prostacyclins, 323 prostaglandin(s), 323, 324 protease inhibitors, 314 proteases, xi, xii, 100, 102, 103, 113, 115, 136, 142, 149, 209, 210, 211, 212, 215, 220, 223, 225, 227, 260, 265, 273 protein design, 192 protein engineering, 2, 193, 225, 329 protein folding, xi, 210, 211, 215 protein function, 192 protein immobilization, 68 protein structure, 5, 160, 192 protein synthesis, 111, 116 proteinase, 103, 212, 265, 276, 280 proteolysis, 80, 81 proteolytic enzyme, 225, 260, 265 proteomics, 215, 225 protocol(s), 25, 34, 81, 107, 192, 269 protons, 220 Prozac, 313 Pseudomonas, 75, 101, 121, 141, 145, 162, 167, 168, 191, 205, 206, 214, 224, 230, 243, 244, 277, 290, 291, 300, 326 Pseudomonas aeruginosa, 121, 141, 145, 244 psychoactive, 156
Index psychopharmacology, 175 pulmonary hypertension, 313 pulse(s), 121, 270 pure water, 5, 8, 15, 16, 20, 31 purification, x, 48, 49, 51, 52, 64, 72, 81, 82, 87, 106, 130, 140, 148, 150, 178, 195, 199, 206, 217, 218, 223, 230, 255, 261, 267, 268, 271, 272, 276, 278, 279, 283, 301 purines, 73 pyrene, 140 pyrimidine, 49, 53, 54, 57, 58, 59, 60, 61, 62, 63, 65, 66, 67, 69, 70, 76, 77, 83, 86, 89 pyrophosphate, 88
Q quantitative estimation, 19, 283 quartz, 26, 27 quaternary ammonium, 244 quinones, 106, 134
R race, 196, 197, 198, 199, 200, 201, 205, 206, 316, 324, 327, 337, 339, 340 racemates, xiv, 184, 307 racemization, 196, 336, 337, 342 radiation, 21, 210 radius, 282, 293, 294 range, vii, xi, xiv, xv, 24, 25, 34, 35, 52, 54, 79, 86, 100, 103, 126, 127, 133, 136, 147, 164, 165, 173, 209, 213, 214, 215, 219, 220, 223, 234, 242, 270, 308, 309, 313, 319, 322, 327, 336, 337 raw materials, 48, 103, 177 reactants, 55, 202 reaction mechanism, 62, 160 reaction medium, 17, 31, 51, 52, 66, 81, 89, 98, 134, 166, 171, 181, 311 reaction rate, xii, 77, 133, 159, 167, 197, 229, 230, 233, 234, 235, 237, 241, 282, 285, 293 reaction temperature, 67, 133 reaction time, 68, 127, 133, 166, 170, 173, 203 reactive groups, 83 reactive oxygen, 29 reactivity, 65, 169 reading, 212, 268 reagents, 15, 48, 49, 85, 120, 170, 196, 284, 341 real time, 19, 21 recall, 5
361
recognition, 16, 25, 65, 173, 328 recombinant DNA, 3, 231 recombination, ix, 187, 189, 190, 192 recovery, 51, 53, 64, 66, 80, 89, 148, 158, 161, 199, 221, 223, 263, 266, 268, 269, 271, 272, 273 recovery processes, 263 recycling, x, 61, 85, 165, 196, 202, 247 reducing sugars, 116 reduction, viii, xii, 28, 50, 81, 95, 96, 103, 109, 111, 116, 118, 119, 120, 122, 123, 124, 130, 131, 132, 138, 140, 167, 181, 197, 206, 229, 230, 239, 268, 271, 294, 309, 311, 312, 313, 314, 315, 316, 318, 339, 340 refining, 153, 173 reflectivity, 21 refractive index, 21 regeneration, xv, 231, 290, 308, 309, 310, 319 regioselectivity, vii, xv, 75, 76, 78, 80, 81, 85, 308, 329 regulation, 48, 138, 191, 262 regulators, 214, 275 regulatory requirements, 51 relationship(s), ix, 34, 103, 116, 162, 179, 187, 213, 225, 340 relevance, 52, 67 remediation, 109, 191 reparation, 332 replication, 188 repression, 101, 102, 137, 148 reserves, 158 residues, viii, 34, 48, 51, 53, 55, 56, 60, 62, 63, 95, 96, 97, 98, 101, 105, 108, 111, 112, 113, 115, 116, 118, 121, 133, 136, 148, 150, 171, 189, 279, 285 resins, 68 resistance, vii, x, 1, 3, 68, 114, 115, 130, 163, 195, 199, 211, 216, 233, 239, 283, 284, 285, 292, 293, 299 resolution, vii, x, xiv, 2, 4, 181, 184, 195, 196, 197, 198, 199, 200, 201, 202, 203, 204, 205, 206, 207, 308, 316, 327, 328, 336, 339, 340, 341, 342 resonator, 27 resources, ix, 147, 155, 160 respiration, 116, 139 restaurants, 121, 122 restructuring, 168 retardation, 295 retention, 63, 115, 117, 125, 139, 283, 284, 285, 327 reverse transcriptase, 188 ribonucleic acid, 266
Index
362 ribose, 50, 53, 55, 60, 63, 72, 81, 83 rice, 97, 98, 101, 135 rice husk, 135 rigidity, 81, 169, 235 rings, 266 Rio de Janeiro, 95, 124, 138 risk, 52, 68 RNA, viii, 24, 47, 84, 188, 266 robotics, 266 rolling, 188 Rome, 142 room temperature, 2, 127, 174, 247 roughness, 250, 255 Russia, 214
S Saccharomyces cerevisiae, 117, 118, 199, 206 safety, xiv, 48, 84, 97, 204, 307 sales, xiv, 301, 307 salinity, xiii, 210, 211, 215, 220, 259 Salmonella, 243, 337 salt(s), xi, xii, xiii, 11, 16, 23, 59, 68, 130, 209, 210, 211, 214, 217, 218, 219, 220, 221, 223, 225, 259, 260, 261 sample, 13, 17, 18, 26, 111, 269, 270 saturated fatty acids, 120 saturation, 28, 31, 164, 167, 189, 192 scalability, 329 scaling, 80 scanning electron microscopy, 28 scattered light, 34 scattering, 6, 21, 253 schizophrenia, 175 science, 292 scientific community, 223 search, vii, 1, 3, 64, 101, 113, 173, 193, 211, 222 seawater, 211, 218, 267, 274, 275 second generation, 265 secrete, 210, 212, 223, 224 secretion, xi, 151, 209, 210, 233, 244 sedimentation, 120, 127 sediments, 212, 222, 227, 261, 269, 278 seed(s), 100, 103, 105, 110, 140, 144, 145, 150, 151, 157, 158, 163, 165, 166, 172, 173, 176, 180, 182, 332, 343 selecting, 80, 288 selectivity, vii, viii, ix, xiv, 2, 47, 49, 51, 52, 63, 72, 76, 81, 82, 85, 86, 89, 155, 161, 163, 174, 183, 191, 221, 308, 329, 340
self-assembly, 35 semiconductors, 267 sensing, 262, 263, 275 sensitivity, xi, 209, 210 separation, ix, 35, 52, 79, 87, 130, 139, 156, 158, 161, 164, 165, 166, 177, 197, 202, 203, 251, 252, 268, 270, 272, 336 sequencing, xi, xii, 119, 121, 124, 138, 190, 209, 212, 213, 214, 217, 224, 230, 231, 265, 266 sequencing batch reactors, 119 series, 106, 130, 156, 160, 163, 303, 309, 314 serine, 103, 115, 138, 160, 162, 179, 212, 216, 217, 223, 233, 265, 273, 322 serotonin, 156, 266, 313 serum, 29, 30, 79, 262 serum albumin, 29, 30, 262 sewage, 145 shape, 4, 19, 20, 25, 52, 57, 81, 103, 166, 289 shear, 27, 269, 286, 289, 299 sheep, 113 shock, 128, 141, 286, 290 shortage, 102 sign, 10, 251 signal transduction, 262, 274, 275 signaling, 263, 275, 285 signals, 275, 285 signal-to-noise ratio, 21 silica, 24, 29, 31, 34, 267, 283 silicate, 267, 269 silicon, 26 similarity, 55, 224 simulation, 147 Singapore, 187 sinus, 324 sites, 55, 58, 109, 132, 180, 192, 213, 285, 290 skin, 324 sludge, 119, 120, 121, 123, 125, 126, 127, 128, 129, 130, 141, 143, 144, 149, 150, 286 small intestine, 171 smoking, 322 smoking cessation, 322 snakes, 260 soda lakes, xi, 209 sodium, xiii, 245, 246, 272 soil, 121, 141, 146, 222, 224, 225, 226, 227, 267, 331 solid matrix, 52, 99 solid state, ix, 8, 95, 96, 97, 103, 105, 108, 111, 115, 116, 117, 118, 124, 143, 145, 146, 147, 148, 149, 150, 152, 153
Index solid surfaces, 283 solid waste, 115, 118, 123, 144, 147, 149 solubility, xiii, 85, 86, 133, 164, 165, 172, 245, 246, 250, 253, 272 solvent(s), ix, 23, 31, 33, 49, 50, 52, 76, 87, 90, 145, 156, 161, 164, 165, 167, 169, 174, 178, 181, 184, 198, 214, 215, 216, 217, 218, 231, 252, 323 soybean, 101, 104, 149, 150, 162, 168, 256 space environment, 299 space-time, 166 Spain, 245 species, xiv, 24, 25, 27, 28, 31, 35, 54, 55, 57, 61, 89, 101, 118, 121, 132, 151, 156, 158, 206, 210, 216, 217, 221, 222, 227, 246, 260, 262, 263, 264, 265, 275, 285, 288, 289, 293, 308 specificity, ix, xiv, 51, 54, 55, 60, 61, 62, 63, 64, 65, 69, 73, 74, 85, 86, 87, 88, 89, 100, 130, 131, 155, 162, 163, 167, 169, 170, 188, 191, 192, 243, 276, 308, 324, 343 spectrophotometry, 243 spectroscopy, 26 spectrum, xiii, 109, 252, 259, 270 speed, 2, 188, 269 stability, x, xi, xiii, 2, 3, 4, 17, 22, 28, 32, 36, 51, 52, 64, 66, 67, 68, 69, 80, 88, 89, 98, 101, 102, 105, 110, 114, 130, 133, 142, 160, 167, 176, 182, 188, 189, 190, 192, 195, 196, 199, 200, 201, 202, 204, 210, 211, 215, 216, 221, 257, 259, 260, 273, 276, 287, 292, 297, 298, 299 stabilization, 52, 68, 69, 81, 132, 207 stages, ix, 6, 16, 18, 87, 160, 187, 284 standard deviation, 129, 236, 237, 238, 240 standards, 132, 270 starch, 35, 102, 104, 117, 118, 216, 223, 260, 264, 275 starvation, 131, 226 statistical analysis, 190, 191 stereospecificity, x, 63, 73, 196 steroids, 180 sterols, 173, 180 stock, 147, 247 stomatitis, 279 storage, x, 29, 33, 128, 196, 200, 313 strain, x, 55, 101, 105, 111, 113, 114, 115, 116, 117, 122, 123, 124, 145, 150, 188, 195, 198, 199, 212, 213, 214, 215, 216, 217, 218, 222, 224, 226, 234, 235, 237, 239, 241, 242, 261, 263, 271, 275, 276, 292, 311, 320, 324, 341 strain improvement, 188
363
strategies, viii, xiii, 3, 95, 96, 115, 143, 150, 221, 231, 245, 246, 247, 255, 263, 268, 309 strength, 25, 28, 36, 51, 52, 81, 152, 165, 283, 284, 289, 298 Streptomyces avermitilis, 192 stress, 117, 134, 165, 199, 210, 269, 289, 299 structural modifications, 165, 329 structure formation, 18 students, 215 styrene, 342 substrates, ix, xii, xiv, xv, 5, 8, 11, 15, 22, 52, 54, 58, 60, 62, 64, 65, 75, 77, 80, 85, 86, 87, 88, 97, 98, 99, 100, 105, 108, 113, 120, 155, 158, 160, 163, 165, 166, 187, 192, 197, 200, 229, 230, 231, 233, 234, 235, 284, 285, 287, 288, 289, 292, 299, 308, 309, 318, 319, 322, 329, 333 sugar, viii, 35, 47, 49, 50, 55, 60, 62, 63, 64, 65, 70, 72, 75, 82, 84, 85, 86, 89, 116, 322, 331 suicide, 107, 142 sulfate, 271, 273, 279, 327 sulfur, 133, 271 sulfur dioxide, 271 sulphur, 113 supercritical carbon dioxide, 164, 165, 166, 171, 172, 173, 174, 181, 182, 184, 185 supercritical fluids, ix, 155, 160, 165 suppliers, 271 supply, xiii, 281, 287, 292, 294, 299 surface area, 16, 29, 34 surface tension, 5, 8, 9, 15 surfactant(s), 15, 22, 23, 159, 170, 231, 247, 253, 257, 311 survival, 210, 285 suspensions, 80, 81, 270, 335 sustainable development, 96 symbiotic, 261, 264 symbols, 282 symmetry, 57, 158, 288, 293 symptoms, 313 syndrome, 337 synergistic effect, xiv, 307 systems, vii, xv, 1, 3, 4, 9, 15, 29, 34, 36, 37, 99, 109, 119, 120, 121, 126, 127, 128, 130, 132, 134, 135, 139, 143, 144, 147, 149, 150, 152, 161, 169, 174, 185, 214, 215, 221, 225, 242, 256, 260, 262, 263, 271, 274, 275, 292, 304, 308
T tandem mass spectrometry, 222
364
Index
tanks, 97 tannic acid, 106 tannins, 105, 115, 116, 136 Tanzania, 147 targets, 156 technology, ix, 51, 52, 63, 97, 100, 119, 121, 130, 134, 139, 144, 159, 175, 176, 179, 187, 202, 204, 221, 231, 255, 270, 271, 273, 279, 299, 340 temperature, vii, x, xiii, 8, 26, 29, 51, 52, 66, 67, 68, 88, 100, 102, 105, 121, 130, 134, 141, 152, 159, 161, 164, 165, 167, 172, 173, 174, 179, 181, 196, 200, 210, 211, 213, 215, 220, 231, 251, 259, 262, 264, 265, 269, 270, 278, 327 tension, 5, 8 textiles, 223 therapeutic agents, 48, 222 therapeutics, 263 thermal stability, x, 68, 196 thermodynamic, 166 thermophiles, 226 thermophilic, 104, 145, 149, 276 thermostability, x, 189, 195, 199, 210 thin films, viii, 2, 4, 29, 35, 36 thrombin, 279 thymidine, 49, 54, 55, 57, 58, 61, 64, 66, 69, 72, 73, 75, 80, 81, 85, 86 thymine, 57, 67, 70, 74, 79 time, xiii, 12, 16, 21, 25, 26, 27, 30, 31, 35, 73, 100, 102, 107, 111, 116, 117, 118, 121, 124, 125, 130, 134, 191, 201, 202, 203, 235, 238, 259, 260, 262, 269, 282, 288, 293, 296, 332 tissue, 117, 134, 143, 261 titanium, 206, 269 tocopherols, 173 Tokyo, 93, 340 toluene, xii, 188, 191, 214, 217, 229, 231, 233, 234, 235, 236, 244, 284 toluene dioxygenase (TDO), xii, 230, 233, 235, 242 topology, 54 toxic substances, 131 toxicity, 48, 49, 111, 112, 117, 118, 120, 133, 134, 136, 147, 164, 214, 284, 320 toxin, 110, 328 trading, 101 traits, 188, 190, 273 transducer, 6 transesterification, 100, 110, 144, 160, 161, 167, 170, 172, 178, 183, 205, 341 transference, 109, 133
transformation(s), vii, 35, 64, 135, 160, 182, 212, 213, 319, 327, 328, 339, 342, 343 transglutaminase, 24 transition(s), 17, 62, 63, 189, 294 translocation, 244 transmission electron microscopy, 35 transpiration, 130 transport, xiii, 121, 128, 165, 203, 220, 230, 233, 281, 287, 289 transport processes, 287 transportation, 337 transposon, xii, 230, 231, 239, 240, 241 trend, 68, 80, 211, 283, 295 triacylglycerides, 183 triacylglycerol(s), ix, 155, 156, 158, 160, 163, 165, 166, 168, 170, 171, 173, 178, 179 triggers, 253 triglyceride(s), 144, 162, 166, 170, 171, 176, 178, 179, 180, 182, 183 trimer, 62 trypsin, 24 tryptophan, 113, 266 tuberculosis, 226 tumor, 273, 278 tumor growth, 278 type 2 diabetes, 331 tyrosine, 24, 60
U UK, 175 ultrasound, 269 uncertainty, 189, 288 uniform, 25, 26, 27, 286, 289, 298 United States, 97, 137, 144, 191, 302 urea, xi, 29, 35, 157, 209 urease, 5, 23, 29, 30, 33, 35 uridine, viii, 48, 54, 64, 66, 69, 77, 81, 82, 83, 84, 175 urinary tract, 324 UV, 26, 34, 135, 248, 249, 251, 252
V validation, 138, 271 validity, 252 values, 26, 28, 52, 87, 88, 97, 103, 107, 116, 118, 124, 127, 288, 290, 295, 298 van der Waals forces, 285
Index variability, 130, 188 variable(s), 51, 102, 149, 157, 173, 315, 339 variation, xiv, 6, 10, 16, 18, 19, 20, 21, 23, 33, 55, 107, 122, 189, 211, 262, 289, 296, 308 vasodilator, 310 vector, 101, 212 vegetable oil, 158, 159, 168, 172, 174, 178, 179 velocity, 167, 202, 235, 269, 342 Venezuela, 95 versatility, 25, 28, 31, 35, 36, 100, 241 virus(es), 24, 86, 271, 273, 278, 279 viscera, 158 viscosity, 34, 110, 148 visualization, 12, 21, 289 vitamin A, 313, 318 vitamins, 117, 163, 173
W Washington, 191, 276, 300 waste disposal, viii, 95, 96 waste management, 123 waste treatment, 118, 223 wastewater(s), ix, 96, 107, 108, 118, 119, 120, 121, 122, 123, 124, 125, 126, 127, 128, 130, 131, 132, 133, 134, 135, 136, 137, 138, 139, 140, 141, 142, 143, 144, 145, 146, 147, 148, 149, 150, 151, 152, 153, 284, 285, 286, 288, 291, 299 wastewater treatment, 119, 123, 128, 130, 132, 133, 134, 136, 139, 141, 142, 151, 284, 285, 286, 291 weak interaction, 69 wealth, 215 weight loss, 156 West Africa, 117 wheat, 98, 101, 104, 105, 185, 226 wheat germ, 185 whole-cell, xii, xv, 151, 204, 229, 230, 231, 233, 235, 237, 238, 239, 240, 241, 242, 243, 244, 308, 309, 324, 325, 333, 335, 341 wild type, 191, 199, 233, 241 Wilhelmy balance, 8, 9, 10 windows, 12 wine, 286 wood, 132, 144, 151, 224, 283 wool, 113, 122, 138, 139, 140 workers, 24, 54, 79 working conditions, 2, 249 workload, x, 196 World Health Organization (WHO), 117, 142 World War, 97
365
X xenobiotic(s), 285, 286, 291 X-ray diffraction, 256 X-ray photoelectron spectroscopy (XPS), 28, 205 xylenes, 215
Y yeast, 96, 98, 100, 104, 105, 108, 123, 153, 205, 243, 278, 311, 312, 313, 314, 315, 318, 322, 333, 338, 339 yield, vii, xiv, 1, 30, 48, 50, 51, 64, 65, 68, 71, 72, 73, 75, 76, 77, 79, 81, 82, 83, 86, 87, 89, 98, 100, 120, 150, 167, 172, 173, 188, 196, 197, 200, 223, 224, 241, 242, 246, 266, 272, 273, 278, 283, 295, 308, 309, 310, 313, 314, 319, 322, 324, 325, 326, 327, 329, 333, 334, 335, 336, 337
Z zirconium, 269, 283
